Cell-scaffold constructs

ABSTRACT

The present invention relates to the regeneration, reconstruction, repair, augmentation or replacement of organs or tissue structures using scaffolds and autologous cells that are not derived from such organs or tissues.

RELATED APPLICATIONS

This application claims priority to under Section 119(e) and the benefitof U.S. Provisional Application Ser. Nos. 61/111,242 filed on Nov. 4,2008; 61/113,542 filed on Nov. 11, 2008; 61/114,021 filed on Nov. 12,2008; 61/114,382 filed on Nov. 13, 2008; 61/114,388 filed on Nov. 13,2008; 61/201,555 filed on Dec. 10, 2008; 61/201,550 filed on Dec. 10,2008; and 61/201,554 filed on Dec. 10, 2008, the disclosures of whichare incorporated by reference herein in their entirety.

SEQUENCE LISTING

The instant application contains a Sequence Listing which has beensubmitted via EFS-Web and is hereby incorporated by reference in itsentirety. Said ASCII copy, created on Jan. 11, 2010, is namedTGN1014U.txt, and is 3,513 bytes in size.

FIELD OF THE INVENTION

The present invention relates to the regeneration, reconstruction,repair, augmentation or replacement of laminarly organized luminalorgans or tissue structures using scaffolds seeded with cells obtainedfrom autologous sources.

BACKGROUND OF THE INVENTION

Several anomalies can cause the bladder to develop abnormally andrequire surgical augmentation. Conditions such as posterior urethralvalves, bilateral ectopic ureters, bladder extrophy, cloacal extrophy,and spina bifida (ie, myelomeningocele) may cause the bladder to benoncompliant, resulting in a small capacity bladder that generates highpressures. Clinically this causes patients to suffer from incontinencewhile increasing their risk for renal failure due to the high pressuresin the genitourinary system. The current standard of therapy for thesepediatric patients is bladder augmentation through enterocystoplasty(Lewis et al. Br. J. Urol. (1990); 65:488-491). Bladder augmentationinvolves the removal of a section of large bowel from the patient whothen has that tissue connected to the existing bladder to increasecompliance, decrease pressure, and improve capacity. The surgeries arerelatively complex and expensive. Even in patients with a good technicalresult, the procedure is associated with numerous immediate risks andchronic complications. The invasiveness, cost, and complications ofthese surgeries limit their use to only the most severe bladderdeficiencies. A similar surgical procedure is performed in adults whorequire a bladder replacement, many as a result of bladder cancer. Inadults, the entire bladder is resected and replaced with large bowel.Despite the risk of adverse effects, there are approximately 10,000 ofthese procedures performed per year in the United States, includingabout 10% in children with congenital abnormalities and 90% in adultswith acquired disorders such as bladder cancer. There is clearly acompelling medical need for an improved approach that would eliminate orat least substantially reduce the adverse effects associated with thecurrent standard of care.

The human urinary bladder is a musculomembranous sac, situated in theanterior part of the pelvic cavity that serves as a reservoir for urine,which it receives through the ureters and discharges through theurethra. In a human the bladder is found in the pelvis behind the pelvicbone (pubic symphysis) and is above and posteriorly connected to adrainage tube, called the urethra, that exits to the outside of thebody. The urinary bladder is subject to numerous maladies and injurieswhich cause deterioration of the urinary bladder in patients. Forexample, bladder deterioration may result from infectious diseases,neoplasms and developmental abnormalities. Further, bladderdeterioration may also occur as a result of trauma such as, for example,car accidents and sports injury. Urinary diversions are often necessaryin bladder cancer patients. There are over 54,000 new bladder cancercases each year in the United States of America. Most bladder cancersare of epithelial origin, and worldwide, there are approximately 336,000new cases of urothelial carcinomas (transitional cell carcinomas (TCC))annually (Kakizoe (2006) Cancer Sci. 97(9) 821).

Urinary diversion is a way to route and excrete urine from the body whenan individual is unable to urinate due to a damaged or non-functionalurinary system. In general, any condition that blocks the flow of urineand increases pressure in the ureters and/or kidneys may require aurinary diversion. Some common indications for diversion include cancerof the bladder requiring a cystectomy, a neurogenic bladder that impactrenal function, radiation injury to the bladder, intractableincontinence that occurs in women, and chronic pelvic pain syndromes. Ingeneral, two major strategies exist for urinary diversion: a urostomyand a continent diversion. A urostomy involves the creation of a stomain the abdomen which is connected to a conduit inside the body such as ashort segment of the small intestine submucosa (SI) such as the ileum,colon or jejunum. In this procedure, the other end of the short SI isconnected to the ureters which normally carry urine from the kidney tothe bladder. Urine flows through the ureters into the short SI and outthe stoma to an external collection reservoir. An alternative of thisprocedure is the attach the ureters directly to a stoma, also called aureterostomy. A continent diversion involves the creation of a pouch orreservoir inside the body from a section of the stomach or small orlarge intestine and the use of a stoma may or may not be required. Forexample, a continent cutaneous reservoir may be created by obtaining asegment of the bowel and modifying it into a more spherical shape. Oneend of the modified segment is connected to the ureters and the other toa stoma that leads to an external collection reservoir. Finally, anorthotopic diversion may created by placing the re-shaped segment inplace of the original bladder by connecting one end to the ureters andthe other end to the urethra so the individual may urinate through theurethra instead of through a stoma.

Although small intestinal submucosa (SI) may be used for urinarydiversion, it has been reported that the removal of the mucosa andsubmucosa may lead to retraction of the intestinal segment (see, e.g.,Atala, A., J. Urol. 156:338 (1996)). Other problems have been reportedwith the use of certain gastrointestinal segments for bladder surgeryincluding stone formation, increased mucus production, neoplasia,infection, metabolic disturbances, long term contracture and resorption.The use of natural materials for urinary diversion has shown thatbladder tissue, with its specific muscular elastic properties andurothelial impermeability functions, cannot be easily replaced. Inaddition, the use of a patient's own bowel segments for urinarydiversion requires at least two different surgical procedures where afirst surgery is performed to remove a segment and a second surgery toinstall the urinary diversion. The requirement of multiple surgeriesincreases the overall cost of the procedures, the risk to the patient,and patient's overall comfort.

Therefore, due to the multiple complications associated with the use ofgastrointestinal segments for urinary diversion and requirement formultiple surgical procedures, there exists a need for methods anddevices for providing urinary diversion systems to patients in need ofsuch a system.

Urinary incontinence is a prevalent problem that affects people of allages and levels of physical health, both in the community at large andin healthcare settings. Medically, urinary incontinence predisposes apatient to urinary tract infections, pressure ulcers, perineal rashes,and urosepsis. Socially and psychologically, urinary incontinence isassociated with embarrassment, social stigmatization, depression, andespecially for the elderly, an increased risk of institutionalization(Herzo et al., Ann. Rev. Gerontol. Geriatrics, 9:74 (1989)).Economically, the costs are astounding; in the United States alone, overten billion dollars per year is spent managing incontinence.

Incontinence can be attributed to genuine urinary stress (bladder andurethra hypermobility), to intrinsic sphincter deficiency (“ISD”), orboth. It is especially prevalent in women, and to a lesser extentincontinence is present in children (in particular, ISD), and in menfollowing radical prostatectomy.

Stress incontinence is an involuntary loss of urine that occurs duringphysical activities which increase intra-abdominal pressure, such ascoughing, sneezing, laughing, or exercise. A person can suffer from oneor both types of incontinence, and when suffering from both, it iscalled mixed incontinence. Despite all of the knowledge associated withincontinence, the majority of cases of urge incontinence are idiopathic,which means a specific cause cannot be identified. Urge incontinence mayoccur in anyone at any age, and it is more common in women and theelderly.

The detrusor is the bladder wall muscle that contracts to expel theurine from the bladder. Consequences of detrusor malfunction such ashyperreflexia include poor bladder compliance, high intravesicalpressure, and reduction in bladder capacity, all of which may result indeterioration of the upper urinary tract.

One current treatment for urge incontinence is injection of neurotoxins,such as botulinum toxin, e.g., Botox®. It is thought that botulinumtoxin exerts its effect on bladder hyperactivity by paralyzing thedetrusor muscle in the bladder wall or possibly impacting afferentpathways in the bladder and reducing sensory receptors in suburothelialnerves. The large size of the botulinum toxin molecule can limit itsability to diffuse, and thus prohibits it from reaching both afferentand efferent nerve fibers. As a result, current methods ofadministration for overactive bladder (OAB), for example, require manyinjections (typically 20 to 50) of botulinum toxin into the bladdermuscle wall, thus increasing the number of doctor visits and associatedcost of treatment. Moreover, the safety of chronic long-term impact ofinhibition of sensory neurotransmitter release from bladder has not yetbeen determined.

Further approaches for treatment of urinary incontinence involveadministration of drugs with bladder relaxant properties, withanticholinergic medications representing the mainstay of such drugs. Forexample, anticholinergics such as propantheline bromide, and combinationsmooth muscle relaxant/anticholinergics such as racemic oxybutynin anddicyclomin, have been used to treat urge incontinence. (See, e.g., A. J.Wein, Urol. Clin. N. Am., 22:557 (1995)). Often, however, such drugtherapies do not achieve complete success with all classes ofincontinent patients, and often results in the patient experiencingsignificant side effects.

Besides drug therapies, other options used by the skilled artisan priorto the present invention include the use of artificial sphincters (LimaS. V. C. et al., J. Urology, 156:622-624 (1996), Levesque P. E. et al.,J. Urology, 156:625-628 (1996)), bladder neck support prosthesis (KondoA. et al., J. Urology, 157:824-827 (1996)), injection of cross-linkedcollagen (Berman C. J. et al., J. Urology, 157:122-124 (1997), Perez L.M. et al., J. Urology, 156:633-636 (1996); Leonard M. P. et al., J.Urology, 156:637-640 (1996)), and injection of polytetrafluoroethylene(Perez L. M. et al., J. Urology, 156:633-636 (1996)).

A recent well known approach for the treatment of urinary incontinenceassociated with ISD is to subject the patient to periurethral endoscopiccollagen injections. This augments the bladder muscle in an effort toreduce the likelihood of bladder leakage or stress incontinence.

Existing solutions to circumvent incontinence have well known drawbacks.While endoscopically directed injections of collagen around the bladderneck has a quite high success rate in sphincter deficiency with nosignificant morbidity, the use of collagen can result in failures thatoccur after an average of two years and considerations need to be givento its cost effectiveness (Khullar V. et al., British J. Obstetrics &Gynecology, 104:96-99 (1996)). In addition, deterioration of patientcontinency, probably due to the migration phenomena (Perez L. M. et al.)may require repeated injections in order to restore continency(Herschorn S. et al., J. Urology, 156:1305-1309 (1996)).

The results with using collagen following radical prostatectomy for thetreatment of stress urinary incontinence have also been generallydisappointing (Klutke C. G. et al., J. Urology, 156:1703-1706 (1996)).Moreover, one study provides evidence that the injection of bovinedermal collagen produced specific antibodies of IgG and IgA class.(McCell and, M. and Delustro, F., J. Urology 155, 2068-2073 (1996)).Thus, possible patient sensitization to the collagen could be expectedover the time.

Despite of the limited success rate, transurethral collagen injectiontherapy remains an acceptable treatment for intrinsic sphincterdeficiency, due to the lack other suitable alternatives.

At present, individuals who suffer from Overactive Bladder Disorders orUrge Incontinence are initially treated by physicians with non-invasivepharmaceutical medical products. However, if these non-invasivepharmaceutical products fail, physicians offer a more invasive solution.

Thus, a need exists for a minimally invasive method of enlarging anexisting laminarily organized luminal organ or tissue structure, e.g., abladder.

Tissue engineering principles have been applied to successfully provideimplantable cell-seeded matrices for use in the reconstruction, repair,augmentation or replacement of laminarily organized luminal organs ortissue structures, such as a bladder, a portion of a bladder, or abladder component. As described in Atala U.S. Pat. No. 6,576,019, cellsmay be derived from the patient's own tissue, including the bladder,urethra, ureter, and other urogenital tissue. However, there arechallenges associated with a dependence upon the development andmaintenance of cell culture systems from the primary organ site as thebasic unit for developing new and healthy engineered tissues. Forexample, the treatment of a defective bladder poses a particularchallenge regarding cell sourcing because it stands to reason thatculturing bladder cells from a defective bladder will result in thecultured cells also being defective. Such cells are not a wise choicefor populating an implantable neo-bladder scaffold or matrix. As suchthere is a need for alternative sources of cells that are suitable forseeding on implantable neo-organ/tissue structure scaffold or matrix.

There is a wealth of literature supporting the notion that human adiposetissue is a rich source of adult stem cells (Devlin et al. (2004),Cytotherapy 6:7-14; Awad, et al. (2003), Tissue Engineering 9:1301-12;Erickson et al. (2002), Biochemical and Biophysical ResearchCommunications 290:763-769; Gronthos et al. (2001), Journal of CellularPhysiology 189:54-63; Halvorsen et al. (2001); Metabolism 50:407-413;Halvorsen et al. (2001), Tissue Eng. 6:729-41; Harp et al. (2001),Biochemical and Biophysical Research Communication 281:907-912; Hicok etal. (2004), Tissue Engineering 10:371-380; Safford et al. (2002), Jun.7, 294(2):371-9; Safford et al, (2004), Experimental Neurology187:319-28; Sen et al. (2004), Journal of Cellular Biochemistry81:312-319; Sigal et al. (1994), Hepatology 19:999-1006; Wickham et al.(2003), Clinical Orthopedics and Related Research, 412:196-212; Ashijianet al. (2003), Plast Reconstr Surg. 111:1922-31; De Ugarte et al.(2003), Cells Tissues Organs. 174:101-9; Mizumo et al. (2002), PlastReconstr Surg. 109:199-209; Morizono et al. (2003), Hum Gene Ther.14:59-66; Winter et al. (2003), Arthritis Rheum. 48:418-29; Zuk et al.(2001), Tissue Eng 7:211-228; Zuk et al. (2002), Mol Biol 13: 4279-4295,reviewed in Gimble et al. (2003), Cytotherapy 5:362-369). These cells,termed Adipose-Derived Adult Stem (ADAS) cells, exhibit animmunophenotype and differentiation potential comparable to that of MSCs(Gronthos et al. (2001), Journal of Cellular Physiology 189:54-63;Safford et al. (2002), Biochem Biophys Res Commun. 294(2):371-9; Zuk etal. (2002), Mol Biol Cell 13:4279-4295).

Reproducible and efficient methods to isolate adult stem cells fromhuman liposuction specimens are available in the public domain (Aust etal. (2004), Cytotherapy 6:7-14; Halvorsen et al. (2001), Metabolism50:407-413). The procedure involves collagenase digestion of the tissue,differential centrifugation, and expansion in culture. A single gram oftissue can yield between 50,000 to 100,000 stromal cells within 24 hoursof culture (Sen et al. (2001), J Cellular Biochemistry 81:312-319).Analysis of specimens obtained from 20 individual donors resulted in aconsistent recovery a mean of 401,000 cells with a viability of 94% froma single ml of liposuction waste (Aust et al. (2004), Cytotherapy6:7-14). Expansion of these cells can result in a population greaterthan 500 million cells within a 2 week period from a standardlipoaspirate.

In the presence of dexamethasone, insulin, isobutylmethylxanthine and athiazolidinedione, the ADAS cells undergo adipogenesis (Sen et al.(2001) Journal of Cellular Biochemistry 81:312-319). The differentiationpotential of the ADAS cells is not limited to the adipocyte lineage.Conditions that promote ADAS cell differentiation along the chondrocyteand osteoblast pathways have been reported (Awad, et al. (2003), TissueEngineering 9:1301-12; Erickson et al. (2002), Biochemical andBiophysical Research Communications 290:763-769; Halvorson et al.(2001), Metabolism 50:407-413; Hicok et al. (2004), Tissue Engineering10:371-380; Wickham et al. (2003), Clinic Orthopedics and RelatedResearch 412:196-212). In vivo, human ADAS cells combined with ahydroxyapatite biomaterial synthesize osteoid matrix when implantedsubcutaneously into immunodeficient mice (Hikok et al. (2004), TissueEngineering 10:371-380). Substantial data are available to demonstratethat murine or human adipose derived adult stem cells (muADAS and huADASrespectively) cultured in the presence of antioxidants and othermediators undergo morphologic and phenotypic changes consistent withneuronal differentiation (De Ugarte (2003) Cells Tissues Organs.174:101-9; Safford et al. (2002), 294(2):371-9; Safford et al. (2004),Experimental Neurology 187:319-28).

As described by Jayo et al. Regen. Med. (2008) 3(5), 671-682(hereinafter referred to as “Jayo I”), attempts to repair organs ortissue have been characterized by incomplete tissue replacementfrequently with collagen deposition, and in some cases scar tissueformation. Jayo et al. also observed a more desirable outcome of tissueengineering is regeneration of the original structure and function of atissue structure or organ. See also Jayo et al., J. Urol. (2008) 180;392-397 (hereinafter referred to as “Jayo II”). Certain molecules arebelieved to be associated with the regenerative process in vivo. Forexample, the chemokine MCP-1 is best known for its ability to recruitmononuclear cells. However, it also appears to be a potent mitogen forvascular smooth muscle cell proliferation. MCP-1 recruits circulatingmonocytes to the area of vessel injury, which in turn are typicallytransformed to macrophages that can serve as reservoirs for cytokinesand growth factors. Macrophages also ingest cholesterol and oxidizelipids. Macrophages and muscle precursor cells are both believed to betargets for MCP-1 signaling. The CCR-2 receptor is the ligand for MCP-1(CCL2) and CCR-2 deficient mice show a regeneration defect with enhancedadipogenesis/fibrosis. Sections from CCR-2 deficient mice whenchallenged with skeletal muscle regeneration demonstrated the followingin comparison to normal mice: more interstitial space, a high number ofinflammatory cells, large round swollen myofibers, more fibroblastaccumulation in interstitial space, fat infiltration with collagendistribution around fat deposits, and fibrosis accompanied by calciumdeposition (Warren et al. (2005), FASEB J. 19:413-415; Selzman et al.(2002), Am J Physiol Heart Circ Physiol. 283(4); H1455-H1461; Shannon etal. (2007), Am. J. Cell Physiol. 292:C953-C967; Shireman et al. (2006),J. Surg. Res. 134(1):145-57. Epub Feb. 20, 2006; Amann et al. (1998),Brit. J. Urol. 82:118-121; Schecter et al. (2004), J. Leukocyte Biol.75:1079-1085; Deonarine et al.,(2007), Transl Med. 5:11; Lumeng et al.(2007), J Clin. Invest. 117(1): 175-184).

The present invention concerns cell populations derived from autologoussources that are different from the organ or tissue structure that isthe subject of the regeneration, reconstruction, repair, augmentation orreplacement described herein, methods of isolating such cells,neo-organ/tissue structure scaffolds or matrices seeded with such cells(constructs) and methods of making the same, as well as methods oftreating a patient in need using such neo-organ/tissue structureconstructs.

SUMMARY OF THE INVENTION

The present invention relates to the regeneration, reconstruction,repair, augmentation or replacement of laminarly organized luminalorgans or tissue structures in a subject in need using scaffolds seededwith autologous cells derived from the subject.

In one aspect, the present invention provides urinary diversionconstructs and methods of making and using the same. In one embodiment,the urinary diversion is for a defective bladder in a subject andincludes (a) a first implantable, biocompatible construct comprising atubular scaffold having a first end configured to connect to anabdominal wall section, a second closed end, and at least a first sideopening configured to connect to a first ureter; and (b) an autologouscell population that is not derived from the defective bladder,deposited on or in a surface of the scaffold. In another embodiment, theurinary diversion is for a defective bladder in a subject and includes(a) an implantable, biocompatible tubular scaffold adapted for temporarystorage and passage of urine that comprises a first end configured toconnect to an opening in the subject's abdominal wall, a second closedend, and at least a first side opening adapted to connect to a firstureter to allow passage of urine from the first ureter to the interiorof the tubular scaffold; and (b) an autologous cell population that isnot derived from the defective bladder, deposited on or in a surface ofthe scaffold.

In one embodiment, the present invention provides a method of preparinga urinary diversion construct for a defective bladder in a subject inneed that includes the steps of a) providing a first implantablebiocompatible scaffold comprising a tubular scaffold having a first endconfigured to contact an abdominal wall section, a second closed end,and at least a first side opening configured to connect to a firstureter; and b) depositing an autologous cell population that is notderived from the defective bladder on or in a first area of the scaffoldto form a urinary diversion construct. In another embodiment, the methodincludes the steps of a) providing an implantable, biocompatible tubularscaffold adapted for temporary storage and passage of urine thatcomprises a first end configured to connect to an opening in thesubject's abdominal wall, a second closed end, and at least a first sideopening adapted to connect to a first ureter to allow passage of urinefrom the first ureter to the interior of the tubular scaffold; and b)depositing an autologous cell population that is not derived from thedefective bladder on or in a surface of the scaffold to form a urinarydiversion construct.

In one other embodiment, the present invention provides a method ofproviding a urinary diversion for a defective bladder in a subject inneed that includes the steps of a) providing a first implantablebiocompatible scaffold comprising a tubular scaffold having a first endconfigured to connect to an abdominal wall section, a second closed end,and at least a first side opening configured to connect to a firstureter; and b) depositing an autologous cell population that is notderived from the defective bladder on or in a first area of the scaffoldto form a urinary diversion construct; and c) implanting the constructinto the subject for the formation of the urinary diversion. In anotherembodiment, the method includes the steps of a) providing animplantable, biocompatible tubular scaffold adapted for temporarystorage and passage of urine that comprises a first end configured toconnect to an opening in the subject's abdominal wall, a second closedend, and at least a first side opening adapted to connect to a firstureter to allow passage of urine from the first ureter to the interiorof the tubular scaffold; b) depositing an autologous cell populationthat is not derived from the defective bladder on or in a surface of thescaffold to form a urinary diversion construct; and c) implanting theconstruct into the subject for the formation of the urinary diversion.In one other embodiment, the method includes the step of implanting intothe subject a urinary diversion construct comprising (a) a tubularscaffold having a first end configured to contact an abdominal wallsection, a second closed end, and at least a first side openingconfigured to connect to a first ureter; and (b) an autologous cellpopulation that is not derived from the defective bladder, deposited onor in a surface of the scaffold, for the formation of the urinarydiversion.

In all embodiments, the urinary diversion scaffold may further comprisea second side opening configured to connect to a second ureter. In allembodiments, the first end may be configured to be positioned flush withthe abdominal wall. In all embodiments, the first end may be configuredto be sutured to the skin of the subject. In all embodiments, the firstend may be configured to form a stoma. In all embodiments, the stoma mayfurther comprise a stoma button. In all embodiments, the scaffoldfurther comprises a washer ring configured to form a stoma. In allembodiments, the biocompatible scaffold is biodegradable. In allembodiments, the scaffold may comprise a material selected from thegroup consisting of polyglycolic acid, polylactic acid, and a copolymerof polyglycolic acid and polylactic acid. In all embodiments, the cellpopulation is a smooth muscle cell population. In all embodiments, thediversion may be a replacement for the defective bladder. In allembodiments, the diversion may be temporary. In all embodiments, thediversion may be permanent. In all embodiments, the tubular scaffold mayhave a rectangular cross-section configuration or a triangularcross-section configuration, or a circular cross-section configuration.In all embodiments, the diversion may be free of urothelial cells. Inall embodiments, the methods of the present invention may provide aneo-urinary conduit characterized by urinary-like tissue regeneration.In all embodiments, the regenerated tissue may be characterized by thepresence of one or more of the following: urothelium, lamina propria,and smooth muscle bundles. In all embodiments, the regenerated tissuecan be observed at one or more of the following: ureter-conduit junction(UCJ), cranial portion of the conduit, and mid-atrium portion of theconduit. In all embodiments, the regenerated tissue may be characterizedby the presence of one or more of the following: mucosa, submucosa, andsmooth muscle with a fibrovascular stroma. In all embodiments, theregenerated tissue is continuous urothelium with underlying smoothmuscle. In all embodiments, the urinary conduit forms an epithelializedmucosa upon implantation.

In one aspect, the present invention concerns isolated smooth musclecell populations. In one embodiment, the cell populations are derivedfrom peripheral blood and contain one or more cells having contractilefunction, that are positive for a smooth muscle cell marker. In anotherembodiment, the cell populations are derived from adipose tissue andcontain one or more cells having contractile function that are positivefor a smooth muscle cell marker.

In all embodiments, the cell populations may be characterized by one ormore smooth muscle cell markers selected from the following: myocardin,alpha-smooth muscle actin, calponin, myosin heavy chain, BAALC, desmin,myofibroblast antigen, and SM22. In all embodiments, the cellpopulations may express myocardin (MYOCD). In all embodiments, the term“MYOCD” includes a nucleic acid encoding a MYOCD polypeptide and a MYOCDpolypeptide.

In all embodiments, the contractile function of the cell populations maybe calcium-dependent.

In another aspect, the present invention provides a smooth muscle cell(SMC) population derived directly from human adipose tissue. In oneembodiment, at least one biomarker selected from the group consisting ofOct4B, osteopontin, BMP6, CD44, and IL-1B, GDF5, HGF, LIF, MCAM, RUNX2,VCAM1, PECAM, vWF, and Flk-1 is differentially expressed in the SMCpopulation, relative to its level of expression in human bonemarrow-derived MSCs. In another embodiment, at least one of GDF5, HGF,LIF, MCAM, RUNX2, VCAM1, PECAM, vWF, and Flk-1 is under-expressed in theSMC population, relative to its level of expression in MSCs. In oneother embodiment, at least one of Oct4B, osteopontin, BMP6, CD44, andIL-1B is over-expressed in the SMC population, relative to its level ofexpression in human bone marrow-derived MSCs.

In another embodiment, the present invention provides a smooth musclecell population derived directly from human adipose tissue characterizedby (a) under-expression of at least one of GDF5, HGF, LIF, MCAM, RUNX2,VCAM1, PECAM, vWF, and Flk-1, and (b) over-expression of at least one ofOct4B, osteopontin, BMP6, CD44, and IL-1B, relative to the expressionlevel thereof in human bone marrow-derived MSCs. In yet anotherembodiment, the SMC population is characterized by (a) under-expressionof all of GDF5, HGF, LIF, MCAM, RUNX2, VCAM1, PECAM, vWF, and Flk-1, and(b) over-expression of all of Oct4B, osteopontin, BMP6, CD44, and IL-1B,relative to the expression level thereof in human bone marrow-derivedMSCs.

In one other embodiment, the present invention provides a smooth musclecell population derived directly from adipose tissue that comprises oneor more cells that are CD45+ and/or one or more cells that are CD117+.

In another embodiment, the present invention provides a smooth musclecell population derived directly from human adipose tissue having ashorter proliferative lifespan than human bone marrow-derived MSCs.

In one other embodiment, the present invention provides a smooth musclecell population derived directly from adipose tissue that exhibitscontact-dependant inhibition of proliferation in culture.

In yet another embodiment, the present invention provides a smoothmuscle cell population derived directly from adipose tissuecharacterized by down-regulation of at least one smooth muscle cell(SMC) marker in response to a thromboxane A2 mimetic. In one embodiment,the SMC marker is selected from the group consisting of myocardin andmyosin heavy chain—smooth muscle isoform (SMMHC). In another embodiment,the myocardin and SMMHC are down-regulated in response to a thromboxaneA2 mimetic.

In one embodiment, the smooth muscle cell populations described hereinare purified cell populations.

In one aspect, the present invention provides a preparation orpopulation of cells derived from adipose tissue. In another embodiment,the population is derived from the SVF of adipose tissue. In anotherembodiment, the SVF contains a cell population that is heterogeneous. Inone other embodiment, the population of cells comprises fullydifferentiated smooth muscle cells. In yet another embodiment, thepresent invention provides a population of human adipose-derived smoothmuscle cells that is distinct from a population of human bonemarrow-derived mesenchymal stem cells (MSCs). In one embodiment, thedistinction is based upon transcriptomic, proteomic, and functionalattributes that are different in the human adipose-derived SMCpopulation, as compared to a population of human bone marrow-derivedMSCs. In one embodiment, the cell population is derived from adiposetissue obtained from an autologous source.

In another aspect, the present invention provides methods of isolatingsmooth muscle cell populations from autologous peripheral blood oradipose sources.

In one embodiment, the method includes the steps of (a) contacting aperipheral blood sample with a density gradient material; (b)centrifuging the sample to define a density gradient comprising amononuclear fraction; (c) extracting the mononuclear fraction from thedensity gradient, wherein the fraction contains a cell population havingone or more smooth muscle cells having contractile function that arepositive for a smooth muscle cell marker. In another embodiment, themethod further includes the step of (d) culturing the cell population.In one embodiment, the cultured cell population forms smooth muscle cellcolonies in culture. In yet another embodiment, the colonies form about5 to about 10 days after culture. In a further embodiment, the cellpopulation does not form endothelial colonies. In other embodiments, themethod further comprises expanding the cell population of step (d). Inanother embodiment, the expanded cell population is a purified cellpopulation.

In another embodiment, the method includes the steps of (a) digesting anadipose tissue sample with collagenase; (b) centrifuging the sample todefine a stromal vascular fraction (SVF); and (c) extracting the SVFfrom the sample, wherein the fraction contains a cell population havingone or more smooth muscle cells having contractile function that arepositive for a smooth muscle cell marker. In another embodiment, themethod further includes the step of (d) culturing the cell population.In other embodiments, the method further includes the step of expandingthe cell population from step (d). In one other embodiment, the expandedcell population is a purified cell population.

In a further embodiment, the present invention provides a method ofproviding an isolated smooth muscle cell population that includes thesteps of a) culturing a heterogenous smooth muscle cell preparationderived from a human adipose SVF without the use of smooth muscle celldifferentiation inductive media; and b) isolating a fully differentiatedsmooth muscle cell population from the cultured cell preparation. In oneother embodiment, the culturing step is preceded by the step ofenzymatically digesting adipose tissue. In another embodiment, theculturing step is preceded by the step of centrifuging the digestedadipose tissue to provide the SVF. In other embodiments, the culturingstep is preceded by the step of washing and plating the SVF. In anotherembodiment, the culturing step comprises selecting for cells that areadherent to a cell culture support. In yet another embodiment, theculturing step does not comprise the use of media containing componentsfor inducing smooth muscle cell differentiation. In one otherembodiment, the culturing step comprises the use of cell culture mediacontaining serum, such as fetal bovine serum (FBS), which containsseveral endogenous growth factors. In another embodiment, the culturingstep does not comprise the selection and addition of specific, exogenousgrowth factors to the cell culture media. In general, an “exogenous”growth factor is a growth factor that is selected and added to a cellculture media in addition to the endogenous growth factors that aretypically already provided by the serum component of the media, such asfrom FBS. Exogenous growth factors may be recombinant growth factors. Inone embodiment, the culturing step does not comprise the use of cellculture media containing exogenous growth factors. In anotherembodiment, the culturing step does not comprise the use of cell culturemedia containing recombinant growth factors.

In all embodiments, the smooth muscle cell population is not anadipose-derived stem cell population and/or the smooth muscle cellpopulation is not a mesenchymal stem cell population.

In one other aspect, the present invention provides constructs forproviding a new laminarily organized luminal organ or tissue structureto a subject in need. In one embodiment, the construct includes (a) animplantable construct comprising a polymeric matrix or scaffold; and (b)an autologous cell population deposited on or in a surface of thepolymeric matrix that is not derived from a native organ or tissuecorresponding to the new organ or tissue structure.

In another aspect, the present invention provides constructs forproviding a neo-bladder or portion thereof to a subject in need. In oneembodiment, the construct includes (a) an implantable constructcomprising a polymeric matrix or scaffold; and (b) an autologous cellpopulation that is not derived from the subject's bladder deposited onor in a surface of the polymeric matrix.

In certain embodiments, the shaped polymeric matrix construct has anellipsoid shape. In some embodiments, the shaped polymeric matrixconstruct is formed into a folded configuration at the time ofimplantation. In one embodiment, the shaped polymeric matrix constructis treated prior to the time of implantation such that flexibility ofthe shaped polymeric matrix construct is more flexible at the time ofimplantation. In another embodiment, the shaped polymeric matrixconstruct is about 10 cm in maximal length. In one embodiment, theshaped polymeric matrix construct is about 4 cm in maximal length. Inanother embodiment, the shaped polymeric matrix construct is about 3 cmin maximal length. In yet another embodiment, the shaped polymericmatrix construct is about 4 cm in maximal width. In one embodiment, theshaped polymeric matrix construct has a 2D surface area of about 30 cm².In another embodiment, the shaped polymeric matrix construct has a 2Dsurface area of about 25 cm².

In other embodiments, the constructs contain cell populations having oneor more peripheral blood-derived smooth muscle cells having contractilefunction that are positive for a smooth muscle cell marker, or cellpopulations having one or more adipose tissue-derived smooth musclecells having contractile function that are positive for a smooth musclecell marker. In one embodiment, the cell population of the construct hascalcium-dependent contractile function.

In all embodiments, the construct may be free of other cell populations.In all embodiments, the construct may be free of urothelial cells. Inall embodiments, the autologous cell population deposited on the matrixis a human adipose-derived smooth muscle cell population as describedherein. In all embodiments, the human adipose-derived SMC population maybe derived directly from the SVF and is fully differentiated. In allembodiments, the human adipose-derived SMC population seeded on thematrix may have the capacity to produce MCP-1 upon implantation of theconstruct in the subject at a site in need. In one embodiment, the MCP-1is an attractant for native mesenchymal stem cells to the site ofimplantation.

In another aspect, the present invention provides methods for preparinga new organ or tissue structure construct suitable for implantation intoa subject in need. In one embodiment, the method includes the steps ofa) obtaining a human adipose tissue sample; b) isolating a fullydifferentiated smooth muscle cell population from the sample; c)culturing the cell population; and d) contacting the cell populationwith a shaped polymeric matrix cell construct, wherein steps a), b), c)and d) are performed in about 45 days or less. In all embodiments, thehuman adipose tissue sample is obtained from an autologous source. Inone other embodiment, the method further includes the step of detectingexpression of a smooth muscle cell marker. In another embodiment,expression is mRNA expression. In a further embodiment, the expressionis polypeptide expression. In one embodiment, the polypeptide expressionis detected by intracellular immunoflourescence.

In one other aspect, the present invention provides methods forproviding a laminarily organized luminal organ or tissue structure to asubject in need. In one embodiment, the method includes the steps of a)providing a biocompatible synthetic or natural polymeric matrix shapedto conform to at least a part of the organ or tissue structure in needof the treatment; b) depositing on or in a first area of the polymericmatrix an autologous cell population that is not derived from a nativeorgan or tissue corresponding to the new organ or tissue structure; andc) implanting the shaped polymeric matrix cell construct into thesubject for the formation of a laminarily organized luminal organ ortissue structure. In one other aspect, the present invention providesmethods for providing a neo-bladder or portion thereof to a subject inneed. In one embodiment, the method includes a) providing abiocompatible synthetic or natural polymeric matrix shaped to conform toa bladder or portion thereof; b) depositing an autologous cellpopulation that is not derived from the subject's bladder on or in afirst area of the polymeric matrix; and c) implanting the shapedpolymeric matrix cell construct into the subject for the formation ofthe neo-bladder or portion thereof. In another embodiment, the cellpopulation of step b) of the methods described herein contains one ormore peripheral blood-derived smooth muscle cells having contractilefunction that are positive for a smooth muscle cell marker, or the cellpopulation of step b) contains one or more adipose tissue-derived smoothmuscle cells having contractile function that are positive for a smoothmuscle cell marker. In one other embodiment, the contractile function ofthe cell population is calcium-dependent. In all embodiments, theautologous cell population deposited on the matrix is a humanadipose-derived smooth muscle cell population as described herein. Inall embodiments, the human adipose-derived SMC population seeded on thematrix has the capacity to produce MCP-1 upon implantation of theconstruct in the subject at a site in need. In one embodiment, the MCP-1is an attractant for native mesenchymal stem cells to the site ofimplantation.

In one embodiment, the methods of the present invention further includethe step of wrapping the implanted conduit construct with the subject'somentum, mesentery, muscle fascia, and/or peritoneum to allow forvascularization.

The present invention further relates to the enlargement of laminarlyorganized luminal organs or tissue structures in a subject in need usingscaffolds seeded with autologous cells derived from the subject. In oneaspect, the present invention provides methods of expanding an existinglaminarily organized luminal organ or tissue structure in a subject inneed of such treatment by providing a polymeric matrix or scaffoldshaped to conform to at least a part of the organ or tissue structure inneed of the treatment and of a sufficient size to be laparoscopicallyimplanted, depositing an autologous cell population that is not derivedfrom the organ or tissue structure on or in a first area of thepolymeric matrix, and laparoscopically implanting the shaped polymericmatrix construct into the subject at the site of the treatment such thatthe existing laminarily organized luminal organ or tissue structure isexpanded. In certain embodiments, the luminal organ or tissue structureis a bladder or a part of a bladder.

In another aspect, the instant invention provides methods for increasingbladder volumetric capacity of a bladder in a subject in need of suchtreatment by providing a biocompatible synthetic or natural polymericmatrix shaped to conform to at least a part of the bladder in need ofsuch treatment and of a sufficient size to be laparoscopicallyimplanted, depositing an autologous cell population that is not derivedfrom the subject's bladder on or in a first area of the polymericmatrix, and laparoscopically implanting the shaped polymeric matrixconstruct laparoscopically into the subject at the site of the treatmentsuch that bladder volume capacity is increased. In one embodiment, thebladder volume capacity is increased about 50 mL. In another embodiment,the bladder volume capacity is increased about 100 mL.

In yet another aspect, the present invention provides methods forexpanding a bladder incision site in a bladder of a subject in need ofsuch treatment by providing a biocompatible synthetic or naturalpolymeric matrix shaped to conform to at least a part of the bladder inneed of the treatment and of a sufficient size to be laparoscopicallyimplanted, depositing an autologous cell population that is not derivedfrom the bladder on or in a first area of the polymeric matrix, andlaparoscopically implanting the shaped polymeric matrix constructlaparoscopically into the subject at the site of the treatment such thatthe bladder incision site is expanded.

In still another aspect, the present invention provides methods for thetreatment of urinary incontinence in a subject in need of such treatmentby providing a biocompatible synthetic or natural polymeric matrixshaped to conform to at least a part of the subject's bladder and of asufficient size to be laparoscopically implanted, depositing anautologous cell population that is not derived from the bladder on or ina first area of the polymeric matrix, laparoscopically implanting theshaped polymeric matrix construct laparoscopically into the subject atthe site of the treatment such that bladder volume capacity isincreased.

The present invention further provides kits comprising the polymericmatrix or scaffold of the invention and instructions for use.

In another aspect, the present invention provides a prognostic methodfor monitoring regeneration of a new organ or tissue structure in asubject following implantation. In one embodiment, the method includesthe step of detecting the level of MCP-1 expression in a test sampleobtained from the subject and in a control sample, wherein a higherlevel of expression of MCP-1 in the test sample, as compared to thecontrol sample, is prognostic for regeneration in the subject. Inanother embodiment, wherein the new organ or tissue structure is derivedfrom an autologous smooth muscle cell population described herein. Inone other embodiment, MCP-1 polypeptide expression is detected. Inanother embodiment, MCP-1 polypeptide expression is detected using ananti-MCP-1 agent. In one other embodiment, the anti-MCP-1 agent is anantibody. In another embodiment, MCP-1 polypeptide expression isdetected using immunohistochemistry. In one embodiment, the detectingstep is preceded by the step of obtaining the test sample from thesubject. In another embodiment, the test sample is blood.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1 shows an example of a bladder augmentation scaffold.

FIG. 2 shows an example of a bladder replacement scaffold.

FIG. 3 shows an example of a urinary diversion or conduit scaffold.

FIG. 3A shows an example of a urinary diversion construct havingdifferent types of cross-sectional areas, as well as potential positionsfor openings that may be configured to connect to ureter(s).

FIG. 3B illustrates variations of a urinary diversion construct (A—openclaim ovoid; B—open claim ovoid receptacle; C—closed ovoid receptacleand three tubes).

FIG. 4 shows different applications of a urinary diversion or conduitconstruct.

FIG. 5A-B show examples of a muscle equivalent scaffold.

FIG. 6 depicts images of various muscle equivalent scaffolds in the formof patches or strips.

FIG. 7 depicts different muscle equivalent scaffolds and representativemethods of implantation. FIG. 7 a depicts formation of a flat sheet ofscaffold. FIG. 7 b depicts a laparoscopically-suited scaffold which canbe rolled at the time of implantation and fed through a laparoscopictube and unrolled in the abdominal cavity. FIG. 7 c depicts formation ofa laparoscopically-suited scaffold sheet in a rolled configuration tofacilitate insertion through a laparoscopic tube, after which it isunrolled in the abdominal cavity. FIG. 7 d depicts formation of alaparoscopically-suited scaffold sheet in a folded configuration oraccordion style to facilitate insertion through the tube, after which itis unfolded in the abdominal cavity. FIG. 7 e depicts possible surgicalmethods for the implantation of a muscle equivalent scaffold. FIG. 7 fdepicts implantation sites on an empty and full bladder. FIG. 7 gdepicts a urinary bladder model with surgical slit showing ellipsoidcreated upon sectioning of surface.

FIG. 8 depicts a pre-folded accordion style scaffold sheet to facilitateinsertion through a laparoscope port.

FIG. 9A depicts scaffold pre-cut into strips, then sutured together toallow stacking and insertion into the laparoscope port and secured inplace in the abdominal cavity.

FIG. 9B depicts one scaffold of 18.7 cm in length by 2.0 cm in widthhaving 2 folds.

FIG. 9C depicts one scaffold of 13.3 cm in length by 2.8 cm in widthhaving 3 folds.

FIG. 9D depicts one scaffold of 9.7 cm in length by 4.0 cm in widthhaving 4 folds.

FIG. 9E depicts one scaffold comprised of two pieces, 2 folds each, of9.7 cm in length and 2.0 cm in width.

FIG. 10 shows an example of a configuration for an implanted conduitconstruct.

FIG. 11 shows an example of the implanted components of a temporaryurinary diversion construct.

FIG. 12 shows an example of the implanted components of a permanenturinary diversion construct.

FIG. 13 depicts other applications of the urinary diversion constructs.

FIG. 14 shows cultures of the canine peripheral blood mononuclearfraction (buffy coat).

FIG. 15 shows canine peripheral blood outgrowth cells.

FIG. 16 shows the morphology of canine smooth muscle cells after severalpassages.

FIG. 17 shows smooth muscle cells isolated from porcine and humanadipose.

FIG. 18 shows RT-PCR analysis for gene expression of the smooth musclecell markers.

FIG. 19 shows immunofluorescence protein expression of smooth musclecell markers.

FIG. 20 shows immunostaining of smooth muscle cells isolated from humanperipheral blood.

FIG. 21 shows the results of a contractility assay for porcine smoothmuscle cells isolated from blood (A), adipose tissue (B), and bladdertissue (C).

FIG. 22 shows the growth of smooth muscle cells isolated from humanadipose tissue as a function of the numbers of cells recovered per unitarea.

FIG. 23 shows the growth of smooth muscle cells isolated from porcineadipose (A), peripheral blood (B), and bladder smooth muscle (C) as afunction of the number of recovered cells per passage.

FIG. 24 shows expression of the cytokine MCP-1 for cells isolated fromhuman bladder smooth muscle, adipose, peripheral blood, and bladderurothelium.

FIG. 25 shows the correlation between MCP-1 production and the densityof cells seeded.

FIG. 26 shows ureteral stents.

FIG. 27 shows a neo-conduit construct.

FIG. 28 shows a neo-conduit construct (dashed arrow) attached to ureters(solid arrows).

FIG. 29 shows the inflow end of a construct attached to the ureters(solid arrows) and the outflow end directed towards the surgicallycreated stoma (dashed arrow).

FIG. 30 shows the stoma and catheter for urine drainage.

FIGS. 31-32 show the cytoscopy images of an animal implanted with acell-free scaffold.

FIG. 33 shows the cytoscopy image of an animal implanted with a scaffoldseeded with blood-derived SMCs.

FIGS. 34-36 show the cytoscopy images of an animal implanted with ascaffold seeded with blood-derived SMCs.

FIG. 37 shows the cytoscopy image of an animal implanted with a scaffoldseeded with blood-derived SMCs.

FIGS. 38-39 show the cytoscopy images of an animal implanted with ascaffold seeded with adipose-derived SMCs.

FIGS. 40-42 show the cytoscopy images of an animal implanted with ascaffold seeded with adipose-derived SMCs.

FIG. 43 shows the cytoscopy image of an animal implanted with a scaffoldseeded with adipose-derived SMCs.

FIG. 44 shows a trimming schematic for a urinary conduit.

FIG. 45 shows subgross photographs of animals implanted with aneo-urinary conduit construct.

FIG. 46 shows a photomicrograph of a neo-urinary conduit near theureter-conduit junction in an animal implanted with a neo-urinaryconduit construct.

FIG. 47 shows a photomicrograph of a neo-urinary conduit near theureter-conduit junction in an animal implanted with a neo-urinaryconduit construct.

FIG. 48 shows photomicrographs of the mid-conduit walls of animalsimplanted with a neo-urinary conduit construct.

FIG. 49 shows pyelogram images for an animal implanted with aneo-urinary conduit (bladder SMC scaffold).

FIG. 50 shows pyelogram images for an animal implanted with aneo-urinary conduit (adipose SMC scaffold).

FIG. 51 shows a pyelogram image for an animal implanted with aneo-urinary conduit (adipose SMC scaffold).

FIG. 52 shows a pyelogram image for an animal implanted with aneo-urinary conduit (blood SMC scaffold).

FIG. 53 shows a pyelogram image for an animal implanted with aneo-urinary conduit (blood SMC scaffold).

FIG. 54 shows a pyelogram image for an animal implanted with aneo-urinary conduit (blood SMC scaffold).

FIG. 55 shows pyelogram images for an animal implanted with aneo-urinary conduit (blood SMC scaffold).

FIG. 56 shows pyelogram images for an animal implanted with aneo-urinary conduit (scaffold only).

FIG. 57 shows pyelogram images for an animal implanted with aneo-urinary conduit (scaffold only).

FIG. 58 shows pyelogram images for an animal implanted with aneo-urinary conduit (scaffold only).

FIG. 59 shows a post-fixed conduit (A) and a trimming schematic (B).FIG. 59C identifies areas of the conduit.

FIG. 60 shows a photomicrograph of an animal implanted with aneo-urinary conduit (Adipose SMC scaffold).

FIG. 61 shows a photomicrograph of an animal implanted with aneo-urinary conduit (Bladder SMC scaffold).

FIG. 62 shows a photomicrograph of an animal implanted with aneo-urinary conduit (Adipose SMC scaffold).

FIG. 63 shows a photomicrograph of an animal implanted with aneo-urinary conduit (Bladder SMC scaffold).

FIG. 64 shows histological characteristics of the regenerated urologicaltissue forming the neo-bladder conduit.

FIG. 65 shows the three muscle components of a native uretero-vesicaljunction.

FIG. 66 shows the utero-conduit junction of a recipient of a conduitconstruct.

FIG. 67 shows the histology of an implanted conduit construct.

FIG. 68 shows morphological features of porcine (A) bladder-, (B)adipose-, and (C) peripheral blood-derived smooth muscle cells.

FIG. 69 shows RT-PCR analysis of smooth muscle cell associated markersfrom porcine bladder-, adipose-, and peripheral blood-derived smoothmuscle cells.

FIG. 70 shows immuno-fluorescence analysis of smooth muscle cellassociated markers from porcine bladder, adipose & peripheralblood-derived smooth muscle cells.

FIG. 71 shows contractility of porcine (A) bladder-, (B) adipose-, and(C) peripheral blood-derived smooth muscle cells.

FIG. 72 shows the growth kinetics of (A) bladder-, (B) adipose-, and (C)peripheral blood-derived smooth muscle cells.

FIG. 73 shows the histological characteristics of the regeneratedurological tissue formed from implanted neo-urinary conduit constructs.

FIG. 74 shows the presence of an epithelizlized mucosa in an animalimplanted with a urinary diversion construct.

FIG. 75 shows cystograms for implanted neo-bladder constructs at 4months. A—Bladder-derived SMCs; B—Blood-derived SMCs; C—Adiposetissue-derived SMCs; D—Native bladder baseline.

FIG. 76 shows the (A) capacity and (B) compliance of implantedneo-bladder constructs over time.

FIG. 77 shows the average body weight of the animals at greater than 4months.

FIG. 78 shows the average serum creatinine of the animals at greaterthan 4 months.

FIG. 79 shows the average BUN for the animals at greater than 4 months.

FIG. 80 shows the average alkaline phosphatase (ALP) for the animals atgreater than 4 months.

FIG. 81 shows the total protein average for the animals at greater than4 months.

FIG. 82 shows the white blood cell (WBC) average for the animals atgreater than 4 months.

FIG. 83 shows cystograms for the implanted constructs at greater than 4months (blood-derived SMCs).

FIG. 84 shows cystograms for the implanted constructs at greater than 4months (adipose-derived SMCs).

FIG. 85 demonstrates the role of cycling in human urinary bladderdevelopment.

FIG. 86 demonstrates that bladder capacity increases with age and urineoutput.

FIG. 87A-C demonstrates that cycling influences regenerative outcome.

FIG. 88 depicts the translation of regeneration-enhancing effects ofcycling to clinical outcomes.

FIG. 89 shows the implantation of a muscle equivalent scaffold.

FIG. 90 shows a cystogram of an animal following implantation of amuscle equivalent scaffold at 4 weeks.

FIG. 91 shows expression levels of various markers in adipose-derivedcell populations: (A) adiponection and FABP-4, (B) SMαA, SM22,myocardin, SMMHC; (C) calponin; (D) VECAD, vWF, PECAM, FLT1; and (E) FLKand TEK.

FIG. 92 shows the dependence of smooth muscle markers on media type: (A)SMαA, SM22, myocardin, SMMHC; and (B) calponin.

FIG. 93 shows a comparison of the expression of smooth muscle markerscalponin, myocardin and SMMHC in adipose-derived cells and mesenchymalstem cells.

FIG. 94 shows the expression of SMαA, SM22, myocardin and calponin inadipose-derived cells over time.

FIG. 95 shows the expression of endothelial markers in adipose-derivedcells and mesenchymal stem cells.

FIG. 96 shows the expression of cell surface markers in adipose-derivedcells.

FIG. 97 shows the expression of cell surface markers in mesenchymal stemcells.

FIG. 98 shows a comparative analysis of the proteomic signatures ofMSCs, bladder-derived SMCs, Ad-SMCs, and human aortic smooth musclecells.

FIG. 99 shows the proliferative capacity of adipose-derived cells overtime in culture.

FIG. 100 shows the response of adipose-derived cells and mesenchymalstem cells to U46619.

FIG. 101 shows the results of RT-PCR analysis of mesodermaldifferentiation markers.

FIG. 102 shows the results of RT-PCR analysis of Oct4A/Oct4B expressionin MSC/AdSMC.

DETAILED DESCRIPTION OF THE INVENTION

The present invention concerns cell populations derived from sourcesthat are different from the organ or tissue structure that is thesubject of the reconstruction, repair, augmentation or replacementdescribed herein, methods of isolating such cells, neo-organ/tissuestructure scaffolds or matrices seeded with such cells (constructs) andmethods of making the same, and methods of treating a patient in needusing such neo-organ/tissue structure constructs.

1. Definitions

Unless defined otherwise, technical and scientific terms used hereinhave the same meaning as commonly understood by one of ordinary skill inthe art to which this invention belongs. Principles of TissueEngineering, 3^(rd) Ed. (Edited by R Lanza, R Langer, & J Vacanti), 2007provides one skilled in the art with a general guide to many of theterms used in the present application.

One skilled in the art will recognize many methods and materials similaror equivalent to those described herein, which could be used in thepractice of the present invention. Indeed, the present invention is inno way limited to the methods and materials described. For purposes ofthe present invention, the following terms are defined below.

The term “smooth muscle cell” or “SMC” as used herein refers to acontractile cell that is derived from a source that is different fromthe native organs or tissues that are the subject of the reconstruction,repair, augmentation or replacement constructs and methods as describedherein. The SMCs may be derived from peripheral blood or adipose tissue.For adipose tissue, the SMCs may be derived from a SVF containingvascular tissue. Thus, the SMCs may be derived from the capillaries,arterioles, and venules of the adipose-derived vascular bed, or the SMCsmay be derived from the perivascular niche containing pericytes. Thesmooth muscle cells provided by the present invention, once seeded andcultured on the scaffolds or matrices described herein, are capable offorming the non-striated muscle that is found in the walls of holloworgans (e.g. bladder, abdominal cavity, gastrointestinal tract, etc.)and characterized by the ability to contract and relax. Those ofordinary skill in the art will appreciate other attributes of smoothmuscle cells.

The term “cell population” as used herein refers to a number of cellsobtained by isolation directly from a suitable mammalian tissue sourceand subsequent culturing in vitro. Those of ordinary skill in the artwill appreciate that various methods for isolating and culturing cellpopulations for use with the present invention and the various numbersof cells in a cell population that are suitable for use in the presentinvention and the various numbers of cells in a cell population that aresuitable for use in the present invention. The cell population may be anadipose-derived smooth muscle cell population (SMC) that issubstantially free of adipocytes or non-adherent adipose cells. The SMCpopulation may be characterized by the expression of markers associatedwith smooth muscle cells. The SMC population may also be a purified cellpopulation. The SMC population may be derived from an autologous source.

The term “autologous” refers to derived or transferred from the sameindividual's body. An autologous smooth muscle cell population isderived from the subject who will be recipient of an implantableconstruct as described herein.

The term “marker” or “biomarker” refers generally to a DNA, RNA,protein, carbohydrate, or glycolipid-based molecular marker, theexpression or presence of which in a cultured cell population can bedetected by standard methods (or methods disclosed herein) and isconsistent with one or more cells in the cultured cell population beinga particular type of cell. In general, the term cell “marker” or“biomarker” refers to a molecule expressed in a cell populationdescribed herein that is typically expressed by a native cell. Themarker may be a polypeptide expressed by the cell or an identifiablephysical location on a chromosome, such as a gene, a restrictionendonuclease recognition site or a nucleic acid encoding a polypeptide(e.g., an mRNA) expressed by the native cell. The marker may be anexpressed region of a gene referred to as a “gene expression marker”, orsome segment of DNA with no known coding function.

The term “smooth muscle cell marker” refers to generally to a DNA, RNA,protein, carbohydrate, or glycolipid-based molecular marker, theexpression or presence of which in a cultured cell population can bedetected by standard methods (or methods disclosed herein) and isconsistent with one or more cells in the cultured cell population beinga smooth muscle cell. In general, the term smooth muscle cell (SMC)“marker” or “biomarker” refers to a molecule that is typically expressedby a native smooth muscle cell. The marker may be a polypeptideexpressed by the cell or an identifiable physical location on achromosome, such as a gene, a restriction endonuclease recognition siteor a nucleic acid encoding a polypeptide expressed by the SMC. Themarker may be an expressed region of a gene referred to as a “geneexpression marker”, or some segment of DNA with no known codingfunction. Such markers contemplated by the present invention include,but are not limited to, one or more of the following: myocardin,alpha-smooth muscle actin, calponin, myosin heavy chain, BAALC, desmin,myofibroblast antigen, SM22, and any combination thereof.

The terms “differentially expressed gene,” “differential geneexpression” and their synonyms, which are used interchangeably, refer toa gene whose expression is activated to a higher or lower level in afirst cell or cell population, relative to its expression in a secondcell or cell population. The terms also include genes whose expressionis activated to a higher or lower level at different stages over timeduring passage of the first or second cell in culture. It is alsounderstood that a differentially expressed gene may be either activatedor inhibited at the nucleic acid level or protein level, or may besubject to alternative splicing to result in a different polypeptideproduct. Such differences may be evidenced by a change in mRNA levels,surface expression, secretion or other partitioning of a polypeptide,for example. Differential gene expression may include a comparison ofexpression between two or more genes or their gene products, or acomparison of the ratios of the expression between two or more genes ortheir gene products, or even a comparison of two differently processedproducts of the same gene, which differ between the first cell and thesecond cell. Differential expression includes both quantitative, as wellas qualitative, differences in the temporal or cellular expressionpattern in a gene or its expression products among, for example, thefirst cell and the second cell. For the purpose of this invention,“differential gene expression” is considered to be present when there isan at least about one-fold, at least about 1.5-fold, at least about2-fold, at least about 2.5-fold, at least about 3-fold, at least about3.5 fold, at least about 4-fold, at least about 4.5-fold, at least about5-fold, at least about 5.5-fold, at least about 6-fold, at least about7-fold, at least about 8-fold, at least about 9-fold, at least about10-fold, at least about 10.5-fold, at least about 11-fold, at leastabout 11.5-fold, at least about 12-fold, at least about 12.5-fold, atleast about 13-fold, at least about 13.5-fold, at least about 14-fold,at least about 14.5-fold, or at least about 15-fold difference betweenthe expression of a given gene in the first cell and the second cell, orat different stages over time during passage of the cells in culture.The differential expression of a marker may be in an adipose-derivedcell (the first cell) relative to expression in a mesenchymal stem cellor MSC (the second cell).

The terms “inhibit”, “down-regulate”, “under-express” and “reduce” areused interchangeably and mean that the expression of a gene, or level ofRNA molecules or equivalent RNA molecules encoding one or more proteinsor protein subunits, or activity of one or more proteins or proteinsubunits, is reduced relative to one or more controls, such as, forexample, one or more positive and/or negative controls. Theunder-expression may be in an adipose-derived cell relative toexpression in an MSC.

The term “up-regulate” or “over-express” is used to mean that theexpression of a gene, or level of RNA molecules or equivalent RNAmolecules encoding one or more proteins or protein subunits, or activityof one or more proteins or protein subunits, is elevated relative to oneor more controls, such as, for example, one or more positive and/ornegative controls. The over-expression may be in an adipose-derived cellrelative to expression in an MSC.

The term “contractile function” refers to smooth muscle contractilefunction involving the interaction of sliding actin and myosinfilaments, which is initiated by calcium-activated phosphorylation ofmyosin thus making contraction dependent on intracellular calciumlevels.

The term “contact-dependent inhibition” refers to the halting of cellgrowth when two or more cells come into contact with each other. Theabsence of this property can be observed in cell culture where cellswhose growth is not inhibited by contact can be observed piling on topof each other, similar to foci formation in transformed cell culture.Mesenchymal stem cells do not exhibit this property. In contrast, cellshaving the contact-dependent inhibition property will not be observed topile on top of each other in culture.

The term “peripheral blood” shall generally mean blood circulatingthroughout the body.

The term “adipose tissue” or “fat” shall generally mean loose connectivetissue made up primarily of adipocytes. Adipose tissue can be obtainedfrom various places in the body including, without limitation, beneaththe skin (subcutaneous fat) and around internal organs (visceral fat).

The term “construct” refers to at least one cell population deposited onor in a surface of a scaffold or matrix made up of one or more syntheticor naturally-occurring biocompatible materials. The cell population maybe combined with a scaffold or matrix in vitro or in vivo.

The term “sample” or “patient sample” or “biological sample” shallgenerally mean any biological sample obtained from an individual, bodyfluid, body tissue, cell line, tissue culture, or other source. The termincludes body fluids such as, for example, blood such as peripheralblood or venous blood, urine and other liquid samples of biologicalorigin, such as lipoaspirates, and solid tissue biopsies such as abiopsy specimen (e.g., adipose tissue biopsy), or tissue cultures orcells derived therefrom, and the progeny thereof. The definition alsoincludes samples that have been manipulated in any way after they areobtained from a source, such as by treatment with reagents,solubilization, or enrichment for certain components, such as proteinsor polynucleotides. The definition also encompasses a clinical sample,and also includes cells in culture, cell supernatants, cell lysates,serum, plasma, biological fluid, and tissue samples. The source of asample may be solid tissue, such as from fresh, frozen and/or preservedorgan or tissue sample or biopsy or aspirate; blood or any bloodconstituents; bodily fluids such as cerebral spinal fluid, amnioticfluid, peritoneal fluid, or interstitial fluid; cells from any time inthe development of the subject. The biological sample may containcompounds which are not naturally present with or in the tissue innature such as preservatives, anticoagulants, buffers, fixatives,nutrients, antibiotics, or the like. The sample can be used for adiagnostic or monitoring assay. Methods for obtaining samples frommammals are well known in the art. If the term “sample” is used alone,it shall still mean that the “sample” is a “biological sample” or“patient sample”, i.e., the terms are used interchangeably. A sample mayalso be a test sample.

The term “test sample” refers to a sample from a subject followingimplantation of a construct described herein. The test sample mayoriginate from various sources in the mammalian subject including,without limitation, blood, serum, urine, semen, bone marrow, mucosa,tissue, etc.

The term “control” or “control sample” refers a negative control inwhich a negative result is expected to help correlate a positive resultin the test sample. Alternatively, the control may be a positive controlin which a positive result is expected to help correlate a negativeresult in the test sample. Controls that are suitable for the presentinvention include, without limitation, a sample known to have normallevels of a cytokine, a sample obtained from a mammalian subject knownnot to have been implanted with a construct described herein, and asample obtained from a mammalian subject known to be normal. A controlmay also be a sample obtained from a subject prior to implantation of aconstruct described herein. In addition, the control may be a samplecontaining normal cells that have the same origin as cells contained inthe test sample. Those of skill in the art will appreciate othercontrols suitable for use in the present invention.

The term “patient” refers to any single animal, more preferably a mammal(including such non-human animals as, for example, dogs, cats, horses,rabbits, zoo animals, cows, pigs, sheep, and non-human primates) forwhich treatment is desired. Most preferably, the patient herein is ahuman.

The term “subject” shall mean any single human subject, including apatient, eligible for treatment, who is experiencing or has experiencedone or more signs, symptoms, or other indicators of a deficient organfunction or failure, including a deficient, damaged or non-functionalurinary system. Such subjects include, without limitation, subjects whoare newly diagnosed or previously diagnosed and now experiencing arecurrence or relapse, or are at risk for deficient organ function orfailure, no matter the cause. The subject may have been previouslytreated for a condition associate with deficient organ function orfailure, or not so treated. Subjects may be candidates for a urinarydiversion including, without limitation, subjects having cancer of thebladder requiring a cystectomy, subjects having a neurogenic bladderthat impacts renal function, subjects having radiation injury to thebladder, and subjects having intractable incontinence. The subject maybe newly diagnosed as requiring a urinary diversion, or previouslydiagnosed as requiring a urinary diversion and now experiencingcomplications, or at risk for a deficient, damaged or non-functionalurinary system, no matter the cause. The subject may have beenpreviously treated for a condition associated with a deficient, damagedor non-functional urinary system, or not so treated.

The term “urinary diversion” or “conduit” refers to the resulting organor tissue structure resulting from the subject's interaction over timewith an implanted urinary diversion construct, anastomosed ureters, andadjacent atrium. The atrium is the anterior connecting chamber thatallows for urine passage through the abdominal wall and may be made bythe most anterior tube-like portion of a peritoneal wrap connecting thecaudal end of the construct (located in the intra-abdominal cavity) tothe skin.

The terms “caudal” and “cranial” are descriptive terms relating to theurinary production and flow. The term “caudal” refers to the end of theurinary diversion construct that upon implantation is closest to thestoma, while the term “cranial” refers to the end of the urinarydiversion construct that upon implantation is closest to the kidneys andureters.

The term “detritis” refers to debris formed during the healing andregenerative process that occurs following implantation of a urinarydiversion construct. Detritis can be made up of exfoliated tissue cells,inflammatory exudate and scaffold biodegradation. If the conduit isobstructed (improper outflow) by such debris, then the stagnated debrisforms a detritis or semisolid bolus within the lumen of the conduit.

The term “debridement” refers to surgical or non-surgical removal offoreign matter, or lacerated, devitalized, contaminated or dead tissuefrom a conduit in order to prevent infection, prevent obstruction, andto promote the healing process. The debridement may involved the removalof detritis.

The term “stoma” refers to a surgically created opening used to passurine from the draining outflow end of a urinary diversion construct tooutside the body. The urine is typically collected in a reservoiroutside the body.

The term “stoma port” or “stoma button” refers to means, such as adevice used to maintain the integrity of the stoma opening.

The term “expanding” or “enlarging” as used herein refers to increasingthe size of the existing laminarily organized luminal organ or tissuestructure. For example, in one aspect of the invention, the existinglaminarily organized luminal organ or tissue structure may be enlargedby 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26,27, 28, or 29 percent. In another aspect of the invention, the existinglaminarily organized luminal organ or tissue structure may be enlargedsuch as to increase the existing volumetric capacity of the existinglaminarily organized luminal organ or tissue structure.

The term “volumetric capacity” as used herein refers to the amount ofliquid capable of being contained in a defined area.

“Regeneration prognosis” or “regenerative prognosis” generally refers toa forecast or prediction of the probable course or outcome of theimplantation of a construct described herein. As used herein,regeneration prognosis includes the forecast or prediction of any one ormore of the following: development or improvement of a functionalbladder after bladder replacement or augmentation, development of afunctional urinary diversion after conduit implantation, development ofimproved bladder capacity, and development of improved bladdercompliance. As used herein, “prognostic for regeneration” meansproviding a forecast or prediction of the probable course or outcome ofthe implantation of a new organ or tissue structure. In someembodiments, “prognostic for regeneration” comprises providing theforecast or prediction of (prognostic for) any one or more of thefollowing: development or improvement of a functional bladder afterbladder replacement or augmentation, development of a functional urinarydiversion after conduit implantation, development of bladder capacity orimproved bladder capacity, and development of bladder compliance orimproved bladder compliance.

“Regenerated tissue” refers to the tissue of a new organ or tissuestructure that develops after implantation of a construct as describedherein. The organ or tissue structure may be a bladder or a part of abladder. The regenerated tissue may include a continous urothelium withunderlying smooth muscle.

2. Cell Populations

The present invention provides populations of smooth muscle cells foruse in the reconstruction, repair, augmentation or replacement oflaminarly organized luminal organs or tissue structures in which thecell population comprises at least one cell that has contractilefunction and is positive for one or more smooth muscle cell markers.

As discussed herein, tissue engineering principles have beensuccessfully applied to provide implantable cell-seeded matrices for usein the reconstruction, repair, augmentation or replacement of laminarilyorganized luminal organs and tissue structures, such as a bladder or abladder component, typically composed of urothelial and smooth musclelayers. (Becker et al. Eur. Urol. 51, 1217-1228 (2007); Frimberger etal. Regen. Med. 1, 425-435 (2006); Roth et al. Curr. Urol. Rep. 10,119-125 (2009); Wood et al. Curr. Opin. Urol. 18, 564-569). Smoothmuscle cells may be derived from the patient's own tissue, including thebladder, urethra, ureter and other urogenital tissue. However, there arechallenges associated with dependence upon the development andmaintenance of cell culture systems from the primary organ site as thebasic unit for developing new and healthy engineered tissues, as forexample during treatment of cancerous bladder tissue. Clearly, suchcancerous cells are inappropriate for populating an implantableneo-bladder scaffold or matrix.

The present invention provides cell populations that are derived fromsources that are different from the organ or tissue structure that isthe subject of the reconstruction, repair, augmentation or replacement.In one embodiment, the source is an autologous source.

In another aspect, the cell population expresses markers consistent withor typical of a smooth muscle cell population.

In one other aspect, the present invention provides smooth muscle cellpopulations isolated from sources that are different from the luminalorgan or tissue structure that is the subject of the reconstruction,repair, augmentation or replacement. In a preferred embodiment, theluminal organ or tissue structure is a bladder or portion of a bladder.

In one aspect, the source is peripheral blood. In one embodiment, theperipheral blood-derived smooth muscle cell population is derived from apatient sample. The patient sample may be venous blood.

In one aspect, the source is adipose tissue. In one embodiment, theadipose tissue-derived smooth muscle cell population is derived from apatient sample. The patient sample may be adipose tissue removed duringan abdominalplasty procedure, or lipoaspirates. In a preferredembodiment, the patient sample

In yet one other embodiment, the isolated cell populations of thepresent invention, upon culturing, can develop various smooth musclecell characteristics including, but not limited to, hill-and-valleymorphology, expression of one or more smooth muscle cell markers,contractile function, filament formation, and cytokine synthesis.

In one aspect, the cultured cell population is characterized by itshill-and-valley morphology. The cells having a hill-and-valleymorphology may have various characteristics including, withoutlimitation, spindly shaped, flattened and fibroblast-like upon passage,elongated and arranged in parallel rows, a “whirled” appearance ofgrowth, and any combination thereof. In one embodiment, the cellpopulation upon culturing in the appropriate media develops a“hill-and-valley morphology” that is typical of cultured smooth musclecells.

In another aspect, the cultured cell population is characterized by thepresence of one or more smooth muscle cell markers. In one embodiment,the cell population upon culturing in the appropriate media developsdetectable smooth muscle cell markers including, without limitation, oneor more of the following myocardin, alpha-smooth muscle actin, calponin,myosin heavy chain, BAALC, desmin, myofibroblast antigen, SM22, and anycombination thereof.

In another aspect, the cultured cell population is characterized by thepresence of one or more cells that express one or more cell surfacemarkers. In one embodiment, the cell population upon culturing in theappropriate media contains one or more cells that are positive for cellsurface markers including, without limitation, one or more of thefollowing CD73, CD90, CD105, CD166, CD31, CD54, CD56, CD117, and anycombination thereof. In another embodiment, the cell population uponculturing in the appropriate media contains one or more cells that areCD45+, CD31+, CD54+, CD56+, CD90+, and CD105+.

In one other aspect, the cultured cell population is characterized bythe presence of one or more cells having contractile function. In oneembodiment, the cell population upon culturing in the appropriate mediadevelops contractile function. In another embodiment, the contractilefunction is calcium dependent. In one other embodiment, thecalcium-dependent contractile function is demonstrated by inhibition ofcontraction with a calcium chelator. In another embodiment, the calciumchelator is EDTA. Those of ordinary skill in the art will appreciatethat other chelators known in the art may be suitable.

In yet another aspect, the cultured cell population is characterized byfilament formation. In one embodiment, the cell population uponculturing in the appropriate media undergoes filament formation.

In one aspect, the cell population includes at least one cell expressingone or more cytokines. In one embodiment, the cytokine is selected fromthe group consisting of MCP-1, oncostatin M, IL-8, and GRO.

In one aspect, the cell populations of the present invention have afinite proliferative lifespan in culture following isolation. In otherembodiments, the cell population has a lifespan of about 1 passage,about 2 passages, about 3 passages, about 4 passages, about 5 passages,about 6 passages, about 7 passages, about 8 passages, about 9 passages,about 10 passages, about 11 passages, about 12 passages, about 13passages, about 14 passages, about 15 passages, about 16 passages, about17 passages, or about 18 passages. In a preferred embodiment, the cellpopulation has a lifespan in culture of no more than 5 passages. Theadipose-derived SMCs can generally be cultured 3-5 days between passagesand the blood-derived SMCs can generally be cultured 14 days before thefirst passage and then 3-5 days for additional passages (see Example 1for more details)

In one aspect, the present invention provides a regenerative cellpopulation containing at least one regenerative cell that when depositedon a scaffold or matrix as described herein and implanted into a subjectin need, provides a regenerative effect for the organ or tissuestructure that is the subject of the reconstruction, repair,augmentation, or replacement contemplated herein. A regenerative cellpopulation has the ability to stimulate or initiate regeneration oflaminarly organized luminal organs or tissue structures uponimplantation into a patient in need. In general, the regeneration of anorgan or tissue structure is characterized by the restoration ofcellular components, tissue organization and architecture, function, andregulative development. In addition, a regenerative cell populationminimizes the incompleteness or disorder that tends to occur at theimplantation site of a cell-seeded luminal organ or tissue structureconstruct. Disorganization at the site of implantation can manifestitself as increased collagen deposition and/or scar tissue formation,each of which can be minimized through the use of a regenerative cellpopulation. In addition, certain cellular events are indicative of theregenerative process. In the case of a regenerated bladder or portion ofa bladder using the cell populations and scaffolds described herein, aregenerating organ or tissue structure is composed of a smooth muscleparenchyma with fibrovascular tissue radiating around numerousmicrovessels that extend toward the luminal surface, as well as stromalelements having well developed blood vessels aligned to the mucosalsurface (see Jayo II supra). A regenerating bladder or portion of abladder is also characterized by the presence of spindloid/mesenchymalcells and αSMA positive muscle precursor cells. In one embodiment, theαSMA positive spindloid cells are observed in neostromal tissues andaround multiple neo-vessels (arterioles).

In one embodiment, the present invention provides a cell population thatwhen deposited on a scaffold or matrix as described herein and implantedinto a subject in need, provides a reparative effect for the organ ortissue structure that is the subject of the reconstruction, repair,augmentation, or replacement contemplated herein. In other embodiments,a reparative effect is characterized by scar tissue formation and/orcollagen deposition. Those of skill in the art will appreciate othercharacteristics of repair that are known in the art.

In another aspect, the regenerative cell population provides aregenerative effect characterized by the adaptive regulation of the sizeof a restored laminarly organized luminal organ or tissue structure. Inone embodiment, the regenerative cell population's regenerative effectis the establishment of adaptive regulation that is specific to thesubject that receives the scaffold or matrix seeded with theregenerative cell population. In one embodiment, the adaptive regulationis the replacement or augmentation of a bladder in a subject using aconstruct described herein such that the neo-bladder grows and developsto a size that is proportional to the subject's body size.

In one embodiment, the cell population capable of regenerativestimulation is an MCP-1 producing cell population, which contains atleast one cell that expresses the chemokine product MCP-1. MCP-1regenerative stimulation is characterized by the recruitment of certaincell types to the site of implantation. In one embodiment, MCP-1recruits muscle progenitor cells to the site of implantation toproliferate within the neo-bladder. In another embodiment, MCP-1recruits monocytes to the site of implantation which in turn producevarious cytokines and/or chemokines to facilitate the regenerativeprocess. In one other embodiment, MCP-1 induces omental cells to developinto muscle cells.

In one aspect, the present invention provides the use of specificcytokines, such as MCP-1, as a surrogate marker for tissue regeneration.Such a marker could be used in conjunction with an assessment ofregeneration based on whether function has been reconstituted.Monitoring a surrogate marker over the time course of regeneration mayalso serve as a prognostic indicator of regeneration.

In another embodiment, the cell population is a purified cellpopulation. A purified cell population as described herein ischaracterized by a phenotype based on one or more of morphology, theexpression of markers, and function. The phenotype includes withoutlimitation, one or more of hill-and-valley morphology, expression of oneor more smooth muscle cell markers, expression of cytokines, a finiteproliferative lifespan in culture, contractile function, and ability toinduce filament formation. The phenotype may include other featuresdescribed herein or known to those of ordinary skill in the art. Inanother embodiment, the purified populations are substantiallyhomogeneous for a smooth muscle cell population as described herein. Apurified population that is substantially homogeneous is typically atleast about 90% homogeneous, as judged by one or more of morphology, theexpression of markers, and function. In other embodiments, the purifiedpopulations are at least about 95% homogeneous, at least about 98%homogeneous, or at least about 99.5% homogeneous.

In another embodiment, the smooth muscle cell population is deriveddirectly from human adipose tissue and is characterized by differentialexpression of one or more of the following osteopontin, Oct4B, growthdifferentiation factor 5 (GDF5), hepatocyte growth factor (HGF),leukemia inhibitory factor (LIF), melanoma cell adhesion molecule(MCAM), vascular cell adhesion molecule 1 (VCAM1), PECAM, vWF, Flk-1,runt-related transcription factor 2 (RUNX2), bone morphogenetic protein6 (BMP6), CD44, and IL-1B, relative to its level of expression in humanbone marrow-derived mesenchymal stem cells (MSCs). In one otherembodiment, the SMC population (a) under-expresses one or more of GDF5,HGF, LIF, MCAM, RUNX2, VCAM1, PECAM, vWF, and Flk-1 and/or (b)over-expression one or more of Oct4B, osteopontin, BMP6, CD44, andIL-1B, relative to the expression level thereof in human bonemarrow-derived MSCs. In one other embodiment, the SMC population (a)under-expresses all of GDF5, HGF, LIF, MCAM, RUNX2, VCAM1, PECAM, vWF,and Flk-1 and/or (b) over-expression all of Oct4B, osteopontin, BMP6,CD44, and IL-1B, relative to the expression level thereof in human bonemarrow-derived MSCs.

In another embodiment, the smooth muscle cell population deriveddirectly from adipose tissue that comprises one or more cells that areCD45+ and/or one or more cells that are CD117+.

In other embodiments, the present invention provides a smooth musclecell population derived directly from human adipose tissue having ashorter proliferative lifespan than human bone marrow-derived MSCs. Inanother embodiment, the SMC population exhibits contact-dependantinhibition of proliferation in culture. In one other embodiment, the SMCpopulation derived directly from adipose tissue characterized bydown-regulation of at least one smooth muscle cell (SMC) marker inresponse to a thromboxane A2 mimetic. In other embodiments, the SMCmarker is selected from the group consisting of myocardin and myosinheavy chain—smooth muscle isoform (SMMHC). In another embodiment, themyocardin and SMMHC are down-regulated in response to a thromboxane A2mimetic.

In all embodiments, the SMC population is derived from an autologoussource.

In one aspect, the present invention contemplates the application of thesmooth muscle cell populations described herein for respiratorydisorders. Airway smooth muscle is present in the bronchial tree of mostvertebrates. A respiratory ocular disorder is one in which the subjecthas a defective respiratory system due to improper function of themuscles of the lung. It has been reported that certain cell populationsmay provide beneficial effects when administered to the lung (e.g.,Ohnishi et al. Int J Chron Obstruct Pulmon Dis. December 2008; 3(4):509-514). Individuals with respiratory disorders such as asthma,emphysema, or chronic obstructive pulmonary disease (COPD) could benefitfrom these SMC populations. Individuals with lung cancer could alsobenefit. In one embodiment, an autologous SMC cell population could beisolated from the adipose tissue or peripheral blood of a subject inneed. The cell population could be seeded onto a scaffold suitable forimplantation at a site within the lung of the subject. An advantage ofthe cell populations of the present invention is that suitable SMCs maynot be available for sourcing from the subject's lung if the subject hasa defective respiratory system, e.g., lung cancer. The cell populationsmay be used in cases where part or all of a subject's lung is removed,such as in the case of lung cancer. Upon removal of a lung or a part ofa lung in a subject, an autologous SMC population could be isolated froma biopsy, cultured, seeded on a suitable scaffold, and implanted intothe subject to provide a new lung or new lung tissue structure.

In another aspect, the present invention contemplates the application ofthe SMC populations described herein for ocular disorders. An oculardisorder is one in which the subject has a defective eye due to improperfunction of the muscles of the eye. Smooth muscle is present as ciliarymuscle in the eye and controls the eye's accommodation for viewingobjects at varying distances and regulates the flow of aqueous humourthrough Schlemm's canal. Smooth muscle is also present in then iris ofthe eye. Individuals with ocular disorders such as presbyopia andhyperopia could benefit from these SMC populations. Individuals withlung cancer could also benefit. In one embodiment, an autologous SMCcell population could be isolated from the adipose tissue or peripheralblood of a subject in need. The cell population could be seeded onto ascaffold suitable for implantation at a site within the eye of thesubject. An advantage of the cell populations of the present inventionis that suitable SMCs may not be available for sourcing from thesubject's eye if the subject has a defective eye or due to the limitedavailability of eye tissue. An autologous SMC population could beisolated from a biopsy, cultured, seeded on a suitable scaffold, andimplanted into the subject to provide new eye tissue.

In another embodiment, the smooth muscle cell populations of the presentinvention may be administered to a subject having a respiratory disorderor an ocular disorder without the use of a scaffold, such as byengraftment. Those of ordinary skill in the art will appreciate suitablemethods of engraftment.

In one embodiment, an autologous SMC cell population could be isolatedfrom the adipose tissue or peripheral blood of a subject in need. Thecell population could be seeded onto a scaffold suitable forimplantation at a site within the lung of the subject. An advantage ofthe cell populations of the present invention is that suitable SMCs maynot be available for sourcing from the subject's lung if the subject hasa defective respiratory disorder, e.g., lung cancer. The cellpopulations may be used in cases where part or all of a subject's lungis removed, such as in the case of lung cancer. Upon removal of a lungor a part of a lung in a subject, an autologous SMC population could beisolated from a biopsy, cultured, seeded on a suitable scaffold, andimplanted into the subject to provide a new lung or lung tissuestructure. In another embodiment, the smooth muscle cell populations ofthe present invention may be administered to a subject having arespiratory disorder without the use of a scaffold, such as byengraftment. Those of ordinary skill in the art will appreciate suitablemethods of engraftment.

3. Methods of Isolating Cell Populations

Autologous cell populations are derived directly from the subjects inneed of treatment. The subject's source tissue is generally not the sameas the organ or tissues structure that is in need of the treatment. Anautologous population of cells may be derived from the patient's owntissue such as, for example, from adipose tissue or peripheral blood.The autologous cells may be isolated in biopsies. In addition, the cellsmay be frozen or expanded before use.

To prepare for construction of a cell-seeded scaffold, sample(s)obtained from a subject containing smooth muscle cells are dissociatedinto appropriate cell suspension(s). Methods for the isolation andculture of cells were discussed in issued U.S. Pat. No. 5,567,612 whichis herein specifically incorporated by reference. Dissociation of thecells to the single cell stage is not essential for the initial primaryculture because single cell suspension may be reached after a period,such as, a week, of in vitro culture. Tissue dissociation may beperformed by mechanical and enzymatic disruption of the extracellularmatrix and the intercellular junctions that hold the cells together.Autologous cells can be cultured in vitro, if desired, to increase thenumber of cells available for seeding on scaffold.

Cells may be transfected prior to seeding with genetic material. Smoothmuscle cells could be transfected with specific genes prior to polymerseeding. The cell-polymer construct could carry genetic informationrequired for the long term survival of the host or the tissue engineeredneo-organ.

Cell cultures may be prepared with or without a cell fractionation step.Cell fractionation may be performed using techniques, which is known tothose of skill in the art. Cell fractionation may be performed based oncell size, DNA content, cell surface antigens, and viability. Forexample, smooth muscle cells may be enriched from adipose tissue, whileendothelial cells and adipocytes may be reduced for smooth muscle cellcollection. While cell fractionation may be used, it is not necessaryfor the practice of the invention.

Another optional procedure in the methods described herein iscryopreservation. Cryogenic preservation may be useful, for example, toreduce the need for multiple invasive surgical procedures. Cells takenfrom a biopsy or sample from the subject may be amplified and a portionof the amplified cells may be used and another portion may becryogenically preserved. The ability to amplify and preserve cells mayminimize the number of surgical procedures required. Another example ofthe utility of cryogenic preservation is in tissue banks. Autologouscells may be stored, for example, in a donor tissue bank. As cells areneeded for new organs or tissue structures, the cryopreserved supply ofcells may be used as needed. Patients who have a disease or undergoingtreatment which may endanger their existing organs or tissue structuresmay cryogenically preserve one or more biopsies. Later, if the patient'sown organ or tissue structure fails, the cryogenically preservedautologous cells may be thawed and used for treatment. For example, if acancer reappeared in a new organ or tissue structure after treatment,cryogenically preserved cells may be used for reconstruction of theorgan or tissue structure without the need for additional biopsies.

Smooth muscle cells may be isolated from adipose or peripheral bloodbased on the following general protocols. An adipose biopsy specimen ofsuitable weight (e.g., in grams) and/or area (e.g., cm²) can beobtained. An appropriate volume of peripheral blood (e.g., ml) can beobtained prior to the planned implantation of a new organ or tissuestructure construct.

The following is a representative example of a protocol suitable for theisolation of smooth muscle cells from the stromal vascular fraction(SVF) of adipose, which represents a heterogenous cell populationcomposed of multiple cell types, including endothelial and smooth musclecells as well as cells that are MSC-like as defined by the InternationalSociety for Cellular Therapy (ISCT) criteria (Domini et al. 2006Cytotherapy 8:4, 315-317). A suitable gram weight of adipose tissue(e.g., 7-25 g) can be obtained by biopsy and washed with PBS (e.g., 3times), minced with a scalpel and scissors, transferred into a 50 mLconical tube and incubated at 37° C. for 60 minutes in a solution ofcollagenase (e.g., 0.1 to 0.3%) (Worthington) and 1% BSA in DMEM-HG. Thetubes may be either continually rocked or periodically shaken tofacilitate digestion. The SVF can be pelleted by centrifugation at 600 gfor 10 minutes and resuspended in DMEM-HG+10% FBS. The stromal-vascularfraction may then be used to seed passage zero.

The following is a representative example of a protocol suitable for theisolation of smooth muscle cells from peripheral blood. A suitablevolume of peripheral blood (e.g. 25 ml) may be diluted 1:1 in PBS andlayered with 25 ml Histopaque-1077 (Sigma) in a 50 mL conical tube.Following centrifugation (e.g., 800 g, 30 min), the mononuclear fractioncan be collected, washed once with PBS and resuspended in α-MEM/10% FBS(Invitrogen) to seed passage zero.

Those of ordinary skill in the art will appreciate additional methodsfor the isolation of smooth muscle cells.

In one aspect, the present invention provides methods for isolating anisolated smooth muscle cell population from SVF without the need forconditions that induce differentiation to smooth muscle cells. In oneembodiment, the method comprises a) obtaining adipose tissue, b)digesting the adipose tissue, c) centrifuging the digested adiposetissue to provide a stromal vascular fraction (SVF), d) culturing theSVF without the need for conditions that induce differentiation tosmooth muscle cells, and e) isolating a smooth muscle cell populationfrom the adipose tissue-derived SVF. In one embodiment, the culturingstep comprises washing the SVF, re-suspending the SVF in a cell culturemedia, and plating the re-suspended SVF. In another embodiment, theculturing step comprises providing a cell population that is adherent tothe cell culture support, such as a plate or container. In anotherembodiment, the method further comprises expanding the cultured cellpopulation. In other embodiments, the method further comprises analyzingthe smooth muscle cell population for smooth muscle cellcharacteristics. In one embodiment, the adipose tissue is derived froman autologous source.

In one embodiment, the culturing conditions do not require the use ofcell culture components for inducing differentiation of the adiposetissue SVF-derived cell population to smooth muscle cells. Jack et al.,J Biomaterials 30 (2009) 3529-3270 report that undifferentiated adiposestem cells derived from SVF were incubated in inductive media containingheparin for 6 weeks in order to differentiate the stem cells into smoothmuscle cells (see also Rodriguez U.S. Pat. No. 7,531,355). The stemcells reported by Jack et al. did not require splitting during thisincubation period. In another embodiment, the culturing conditions donot require the use of inductive media, including inductive mediacontaining heparin. In one other embodiment, the methods of the presentinvention comprise the use of culturing conditions that do not requirethe use of exogenous growth factors for differentiating a cellpopulation into smooth muscle cells or for culturing and expanding acell population.

The advantages of the methods of the present invention over otherreported methods include the elimination of the step of differentiatingadipose derived stem cells into smooth muscle cells, which reduces thetime between obtaining an adipose biopsy and isolating a smooth musclecell population therefrom. In addition, the elimination of the need forother cell culture media components for inducing differentiation, suchas exogenous growth factors, is advantageous in terms of cost.

In one other aspect, the present invention provides methods of isolatingand culturing populations of smooth muscle cells that contain at leastone cell that has contractile function and is positive for one or moresmooth muscle cell markers. In one embodiment, the method includes thestep of obtaining a sample from a patient in need of the reconstruction,repair, augmentation or replacement of a laminarily organized luminalorgan or tissue structure, where the sample is not obtained from theluminal organ or tissue structure that is in need of the reconstruction,repair, augmentation or replacement. In another embodiment, smoothmuscle cells are derived from the patient sample. In one otherembodiment, the luminal organ or tissue structure is a bladder orportion of a bladder. In one embodiment, the sample is an autologoussample. In another embodiment, the sample is a peripheral blood sample.In yet another embodiment, the sample is an adipose tissue sample. Theadipose tissue may be tissue removed from a subject as a result of anabdominalplasty procedure.

In another embodiment, the obtaining step is followed by a separationstep.

In the case of a peripheral blood sample, the separation step includescontacting the sample with a density gradient material, centrifuging thesample to define a density gradient that has a mononuclear fraction, andextracting the mononuclear fraction from the density gradient. Theseparation step may be followed by a culturing step in which cells fromthe extracted fraction are cultured.

In the case of an adipose tissue sample, the purification step includesdigestion of the sample with collagenase, centrifuging the digestedsample, mixing of the centrifuged sample to separate stromal cells fromprimary adipocytes, centrifuging the mixed sample to obtain astromal-vascular fraction that can be resuspended for subsequentculturing.

In one aspect, the present invention provides a method of providing anisolated smooth muscle cell (SMC) population without the use ofdifferentiation inductive cell culture media. In one embodiment, themethod includes the steps of a) obtaining an adipose tissue biopsy, b)enzymatically digesting the adipose tissue, c) centrifuging the digestedadipose tissue to provide a stromal vascular fraction (SVF) thatcontains a heterogenous population of cells, d) washing and plating theheterogeneous population of cells; e) culturing the population of cellswithout the use of smooth muscle cell differentiation inductive media,f) isolating a fully differentiated SMC population from the culturedcells. In one other embodiment, the culturing step e) includes selectingfor cells that are adherent to a cell culture support. In anotherembodiment, the culturing step e) does not include the use of cellculture media that contains exogenous growth factors. In one embodiment,the culturing method includes the use of cell culture media containingminimal essential medium (e.g., DMEM or α-MEM) and fetal bovine serum(e.g., 10% FBS) by standard conditions known to those of ordinary skillin the art. In another embodiment, the smooth muscle cell population isnot an adipose-derived stem cell population. In one other embodiment,the smooth muscle cell population is not a mesenchymal stem cellpopulation.

4. Scaffolds

As described in Atala U.S. Pat. No. 6,576,019 (incorporated herein byreference in its entirety), scaffolds or polymeric matrices may becomposed of a variety of different materials. In general, biocompatiblematerial and especially biodegradable material is the preferred materialfor the construction of the scaffolds described herein. The scaffoldsare implantable, biocompatible, synthetic or natural polymeric matriceswith at least two separate surfaces. The scaffolds are shaped to conformto a at least a part of the luminal organ or tissue structure in need ortreatment. The biocompatible materials are biodegradeable. Biocompatiblerefers to materials which do not have toxic or injurious effects onbiological functions. Biodegradable refers to material that can beabsorbed or degraded in a patient's body. Examples of biodegradablematerials include, for example, absorbable sutures. Representativematerials for forming the scaffolds include natural or syntheticpolymers, such as, for example, collagen, poly(alpha esters) such aspoly(lactate acid), poly(glycolic acid), polyorthoesters andpolyanhydrides and their copolymers, which degraded by hydrolysis at acontrolled rate and are reabsorbed. These materials provide the maximumcontrol of degradability, manageability, size and configuration.Preferred biodegradable polymer material include polyglycolic acid andpolyglactin, developed as absorbable synthetic suture material.Polyglycolic acid and polyglactin fibers may be used as supplied by themanufacturer. Other scaffold materials include cellulose ether,cellulose, cellulosic ester, fluorinated polyethylene,poly-4-methylpentene, polyacrylonitrile, polyamide, polyamideimide,polyacrylate, polybenzoxazole, polycarbonate, polycyanoarylether,polyester, polyestercarbonate, polyether, polyetheretherketone,polyetherimide, polyetherketone, polyethersulfone, polyethylene,polyfluoroolefin, polyimide, polyolefin, polyoxadiazole, polyphenyleneoxide, polyphenylene sulfide, polypropylene, polystyrene, polysulfide,polysulfone, polytetrafluoroethylene, polythioether, polytriazole,polyurethane, polyvinyl, polyvinylidene fluoride, regenerated cellulose,silicone, urea-formaldehyde, or copolymers or physical blends of thesematerials. The material may be impregnated with suitable antimicrobialagents and may be colored by a color additive to improve visibility andto aid in surgical procedures.

Other scaffold materials that are biodegradeable include syntheticsuture material manufactured by Ethicon Co. (Ethicon Co., Somerville,N.J.), such as MONOCRYL™ (copolymer of glycolide andepsilon-caprolactone), VICRYL™ or Polyglactin 910 (copolymer of lactideand glycolide coated with Polyglactin 370 and calcium stearate), andPANACRYL™ (copolymer of lactide and glycolide coated with a polymer ofcaprolactone and glycolide). (Craig P. H., Williams J. A., Davis K. W.,et al.: A Biological Comparison of Polyglactin 910 and Polyglycolic AcidSynthetic Absorbable Sutures. Surg. 141; 1010, (1975)) and polyglycolicacid. These materials can be used as supplied by the manufacturer.

In yet another embodiment, the matrix or scaffold can be created usingparts of a natural decellularized organ. Biostructures, or parts oforgans can be decellularized by removing the entire cellular and tissuecontent from the organ. The decellularization process comprises a seriesof sequential extractions. One key feature of this extraction process isthat harsh extraction that may disturb or destroy the complexinfra-structure of the biostructure, be avoided. The first step involvesremoval of cellular debris and solubilization of the cell membrane. Thisis followed by solubilization of the nuclear cytoplasmic components andthe nuclear components.

Preferably, the biostructure, e.g., part of an organ is decellularizedby removing the cell membrane and cellular debris surrounding the partof the organ using gentle mechanical disruption methods. The gentlemechanical disruption methods must be sufficient to disrupt the cellularmembrane. However, the process of decellularization should avoid damageor disturbance of the biostructure's complex infra-structure. Gentlemechanical disruption methods include scraping the surface of the organpart, agitating the organ part, or stirring the organ in a suitablevolume of fluid, e.g., distilled water. In one preferred embodiment, thegentle mechanical disruption method includes stirring the organ part ina suitable volume of distilled water until the cell membrane isdisrupted and the cellular debris has been removed from the organ.

After the cell membrane has been removed, the nuclear and cytoplasmiccomponents of the biostructure are removed. This can be performed bysolubilizing the cellular and nuclear components without disrupting theinfra-structure. To solubilize the nuclear components, non-ionicdetergents or surfactants may be used. Examples of nonionic detergentsor surfactants include, but are not limited to, the Triton series,available from Rohm and Haas of Philadelphia, Pa., which includes TritonX-100, Triton N-101, Triton X-114, Triton X-405, Triton X-705, andTriton DF-16, available commercially from many vendors; the Tweenseries, such as monolaurate (Tween 20), monopalmitate (Tween 40),monooleate (Tween 80), and polyoxethylene-23-lauryl ether (Brij. 35),polyoxyethylene ether W-1 (Polyox), and the like, sodium cholate,deoxycholates, CHAPS, saponin, n-Decyl-D-glucopuranoside,n-heptyl-D-glucopyranoside, n-Octyl-D-glucopyranoside and Nonidet P-40.

One skilled in the art will appreciate that a description of compoundsbelonging to the foregoing classifications, and vendors may becommercially obtained and may be found in “Chemical Classification,Emulsifiers and Detergents”, McCutcheon's, Emulsifiers and Detergents,1986, North American and International Editions, McCutcheon Division, MCPublishing Co., Glen Rock, N.J., U.S.A. and Judith Neugebauer, A Guideto the Properties and Uses of Detergents in Biology and Biochemistry,Calbiochem. R., Hoechst Celanese Corp., 1987. In one preferredembodiment, the non-ionic surfactant is the Triton. series, preferably,Triton X-100.

The concentration of the non-ionic detergent may be altered depending onthe type of biostructure being decellularized. For example, for delicatetissues, e.g., blood vessels, the concentration of the detergent shouldbe decreased. Preferred concentration ranges of non-ionic detergent canbe from about 0.001 to about 2.0% (w/v). More preferably, about 0.05 toabout 1.0% (w/v). Even more preferably, about, 0.1% (w/v) to about 0.8%(w/v). Preferred concentrations of these range from about 0.001 to about0.2% (w/v), with about 0.05 to about 0.1% (w/v) particular preferred.

The cytoskeletal component, which includes the dense cytoplasmicfilament networks, intercellular complexes and apical microcellularstructures, may be solubilized using alkaline solution, such as,ammonium hydroxide. Other alkaline solution consisting of ammonium saltsor their derivatives may also be used to solubilize the cytoskeletalcomponents. Examples of other suitable ammonium solutions includeammonium sulphate, ammonium acetate and ammonium hydroxide. In apreferred embodiment, ammonium hydroxide is used.

The concentration of the alkaline solutions, e.g., ammonium hydroxide,may be altered depending on the type of biostructure beingdecellularized. For example, for delicate tissues, e.g., blood vessels,the concentration of the detergent should be decreased. Preferredconcentrations ranges can be from about 0.001 to about 2.0% (w/v). Morepreferably, about 0.005 to about 0.1% (w/v). Even more preferably,about, 0.01% (w/v) to about 0.08% (w/v).

The decellularized, lyophilized structure may be stored at a suitabletemperature until required for use. Prior to use, the decellularizedstructure can be equilibrated in suitable isotonic buffer or cellculture medium. Suitable buffers include, but are not limited to,phosphate buffered saline (PBS), saline, MOPS, HEPES, Hank's BalancedSalt Solution, and the like. Suitable cell culture medium includes, butis not limited to, RPMI 1640, Fisher's, Iscove's, McCoy's, Dulbecco'smedium, and the like.

Still other biocompatible materials that may be used include stainlesssteel, titanium, silicone, gold and silastic.

The polymeric matrix or scaffold can be reinforced. For example,reinforcing materials may be added during the formation of a syntheticmatrix or scaffold or attached to the natural or synthetic matrix priorto implantation. Representative materials for forming the reinforcementinclude natural or synthetic polymers, such as, for example, collagen,poly(alpha esters) such as poly(lactate acid), poly(glycolic acid),polyorthoesters and polyanhydrides and their copolymers, which degradedby hydrolysis at a controlled rate and are reabsorbed. These materialsprovide the maximum control of degradability, manageability, size andconfiguration.

The biodegradable polymers can be characterized with respect tomechanical properties, such as tensile strength using an Instron tester,for polymer molecular weight by gel permeation chromatography (GPC),glass, transition temperature by differential scanning calorimetry (DSC)and bond structure by infrared (IR) spectroscopy; with respect totoxicology by initial screening tests involving Ames assays and in vitroteratogenicity assays and implantation studies in animals forimmunogenicity, inflammation, release and degradation studies. In vitrocell attachment and viability can be assessed using scanning electronmicroscopy, histology and quantitative assessment with radioisotopes.The biodegradable material may also be characterized with respect to theamount of time necessary for the material to degrade when implanted in apatient. By varying the construction, such as, for example, thethickness and mesh size, the biodegradable material may substantiallybiodegrade between about 2 years or about 2 months, preferably betweenabout 18 months and about 4 months, most preferably between about 15months and about 8 months and most preferably between about 12 monthsand about 10 months. If necessary, the biodegradable material may beconstructed so as not to degrade substantially within about 3 years, orabout 4 years or about five or more years.

The polymeric matrix or scaffold may be fabricated with controlled porestructure as described above. The size of the pores may be used todetermine the cell distribution. For example, the pores on the polymericmatrix or scaffold may be large to enable cells to migrate from onesurface to the opposite surface. Alternatively, the pores may be smallsuch that there is fluid communication between the two sides of thepolymeric matrix or scaffold but cells cannot pass through. Suitablepore size to accomplish this objective may be about 0.04 micron to about10 microns in diameter, preferably between about 0.4 micron to about 4microns in diameter. In some embodiments, a surface of the polymericmatrix or scaffold may comprise pores sufficiently large to allowattachment and migration of a cell population into the pores. The poresize may be reduced in the interior of the polymeric matrix or scaffoldto prevent cells from migrating from one side of the polymeric matrix orscaffold to the opposite side. One embodiment of a polymeric matrix orscaffold with reduced pore size is a laminated structure of a small porematerial sandwiched between two large pore material. Polycarbonatemembranes are especially suitable because they can be fabricated in verycontrolled pore sizes such as, for example, about 0.01 microns, about0.05 micron, about 0.1 micron, about 0.2 micron, about 0.45 micron,about 0.6 micron, about 1.0 micron, about 2.0 microns and about 4.0microns. At the submicron level the polymeric matrix or scaffold may beimpermeable to bacteria, viruses and other microbes.

The following characteristics or criteria, among others, are taken intoaccount in the design of each discrete matrix, or part thereof: (i)shape, (ii) strength, (iii) stiffness and rigidity, and (iv)suturability (the degree to which the matrix, or part thereof, isreadily sutured or otherwise attached to adjacent tissue). As usedherein, the stiffness of a given matrix or scaffold is defined by themodulus of elasticity, a coefficient expressing the ratio between stressper unit area acting to deform the scaffold and the amount ofdeformation that results from it. (See e.g., Handbook of Biomaterialsevaluation, Scientific, Technical, and Clinical Testing of ImplantMaterials, 2nd edition, edited by Andreas F. von Recum, (1999); Ratner,et al., Biomaterials Science: An Introduction to Materials in Medicine,Academic Press (1996)). The rigidity of a scaffold refers to the degreeof flexibility (or lack thereof) exhibited by a given scaffold.

Each of these criteria is a variable that can be changed (through, amongother things, the choice of material and the manufacturing process) toallow the matrix, or part thereof to best placed and modified to addressthe medical indication and the physiological function for which it isintended. For example, the material comprising the matrix or scaffoldfor bladder replacement, reconstruction and/or augmentation must besufficiently strong to support sutures without tearing, while beingsufficient compliant so as to accommodate fluctuating volumes of urine.

Optimally, the matrix or scaffold should be shaped such that after itsbiodegradation, the resulting reconstructed bladder is collapsible whenempty in a fashion similar to a natural bladder and the ureters will notbe obstructed while the urinary catheter has been removed from the neworgan or tissue structure without leaving a leak point. Thebioengineered bladder construct can be produced as one piece or eachpart can be individually produced or combinations of the sections can beproduced as specific parts. Each specific matrix or scaffold part may beproduced to have a specific function. Otherwise specific parts may beproduced for manufacturing ease. Specific parts may be constructed ofspecific materials and may be designed to deliver specific properties.Specific part properties may include tensile strength similar to thenative tissue (e.g. ureters) of 0.5 to 1.5 MPa.sup.2 and an ultimateelongation of 30 to 100% or the tensile strength may range from 0.5 to28 MPa.sup.2, ultimate elongations may range from 10-200% andcompression strength may be <12.

A mesh-like structure formed of fibers, which may be round, scalloped,flattened, star shaped, solitary or entwined with other fibers ispreferred. The use of branching fibers is based upon the same principleswhich nature has used to solve the problem of increasing surface areaproportionate to volume increases. All multicellular organisms utilizethis repeating branching structure. Branching systems representcommunication networks between organs, as well as the functional unitsof individual organs. Seeding and implanting this configuration withcells allows implantation of large numbers of cells, each of which isexposed to the environment of the host, providing for free exchange ofnutrients and waste while neovascularization is achieved. The polymericmatrix or scaffold may be made flexible or rigid, depending on thedesired final form, structure and function.

In one preferred embodiment, the polymeric matrix or scaffold is formedwith a polyglycolic acid with an average fiber diameter of 15 .mu.m andconfigured into a bladder shaped mold using 4-0 polyglactin 910 sutures.The resulting structure is coated with a liquefied copolymer, such as,for example, pol-DL-lactide-co-glycolide 50:50, 80 milligram permilliliter methylene chloride, in order to achieve adequate mechanicalcharacteristics and to set its shape.

In a further embodiment, the scaffolds of the present invention arecoated with a biocompatible and biodegradable shape-setting material. Inone embodiment, the shape-setting material contains apoly-lactide-co-glycolide copolymer. In another embodiment, the shapesetting material is liquefied.

In one other aspect, the scaffolds of the present invention may betreated with additives or drugs prior to implantation (before or afterthe polymeric matrix or scaffold is seeded with cells), e.g., to promotethe regeneration of new tissue after implantation. Thus, for example,growth factors, cytokines, extracellular matrix or scaffold components,and other bioactive materials can be added to the polymeric matrix orscaffold to promote graft healing and regeneration of new tissue. Suchadditives will in general be selected according to the tissue or organbeing reconstructed, replaced or augmented, to ensure that appropriatenew tissue is formed in the engrafted organ or tissue (for examples ofsuch additives for use in promoting bone healing, see, e.g.,Kirker-Head, C. A. Vet. Surg. 24 (5): 408-19 (1995)). For example, whenpolymeric matrices (optionally seeded with endothelial cells) are usedto augment vascular tissue, vascular endothelial growth factor (VEGF),(see, e.g., U.S. Pat. No. 5,654,273) can be employed to promote theregeneration of new vascular tissue. Growth factors and other additives(e.g., epidermal growth factor (EGF), heparin-binding epidermal-likegrowth factor (HBGF), fibroblast growth factor (FGF), cytokines, genes,proteins, and the like) can be added in amounts in excess of any amountof such growth factors (if any) which may be produced by the cellsseeded on the polymeric matrix, if added cells are employed. Suchadditives are preferably provided in an amount sufficient to promote theregeneration of new tissue of a type appropriate to the tissue or organ,which is to be repaired, replaced or augmented (e.g., by causing oraccelerating infiltration of host cells into the graft). Other usefuladditives include antibacterial agents such as antibiotics.

One preferred supporting matrix or scaffold is composed of crossingfilaments which can allow cell survival by diffusion of nutrients acrossshort distances once the cell support is implanted. The cell supportmatrix or scaffold becomes vascularized in concert with expansion of thecell mass following implantation.

The building of three-dimensional structure constructs in vitro, priorto implantation, facilitates the eventual terminal differentiation ofthe cells after implantation in vivo, and minimizes the risk of aninflammatory response towards the matrix, thus avoiding graftcontracture and shrinkage.

The polymeric matrix or scaffold may be sterilized using any knownmethod before use. The method used depend on the material used in thepolymeric matrix. Examples of sterilization methods include steam, dryheat, radiation, gases such as ethylene oxide, gas and boiling.

The synthetic materials that make up the scaffolds may be shaped usingmethods such as, for example, solvent casting, compression molding,filament drawing, meshing, leaching, weaving and coating. In solventcasting, a solution of one or more polymers in an appropriate solvent,such as methylene chloride, is cast as a branching pattern reliefstructure. After solvent evaporation, a thin film is obtained. Incompression molding, a polymer is pressed at pressures up to 30,000pounds per square inch into an appropriate pattern. Filament drawinginvolves drawing from the molten polymer and meshing involves forming amesh by compressing fibers into a felt-like material. In leaching, asolution containing two materials is spread into a shape close to thefinal form of the construct. Next a solvent is used to dissolve away oneof the components, resulting in pore formation. (See Mikos, U.S. Pat.No. 5,514,378, hereby incorporated by reference.) In nucleation, thinfilms in the shape of a scaffold are exposed to radioactive fissionproducts that create tracks of radiation damaged material. Next thepolycarbonate sheets are etched with acid or base, turning the tracks ofradiation-damaged material into pores. Finally, a laser may be used toshape and burn individual holes through many materials to form astructure with uniform pore sizes. Coating refers to coating orpermeating a polymeric structure with a material such as, for exampleliquefied copolymers (poly-DL-lactide co-glycolide 50:50 80 mg/mlmethylene chloride) to alter its mechanical properties. Coating may beperformed in one layer, or multiple layers until the desired mechanicalproperties are achieved. These shaping techniques may be employed incombination, for example, a polymeric matrix or scaffold may be weaved,compression molded and glued together. Furthermore different polymericmaterials shaped by different processes may be joined together to form acomposite shape. The composite shape may be a laminar structure. Forexample, a polymeric matrix or scaffold may be attached to one or morepolymeric matrixes to form a multilayer polymeric matrix or scaffoldstructure. The attachment may be performed by gluing with a liquidpolymer or by suturing. In addition, the polymeric matrix or scaffoldmay be formed as a solid block and shaped by laser or other standardmachining techniques to its desired final form. Laser shaping refers tothe process of removing materials using a laser.

In a preferred embodiment, the scaffolds are formed from nonwovenpolygycolic acid (PGA) felts and poly(lactic-co-glycolic acid) polymers(PLGA). In another preferred embodiment, the scaffold is a urinarydiversion scaffold.

As described in Bertram et al. U.S. Published Application 20070276507(incorporated herein by reference in its entirety), the polymeric matrixor scaffold of the present invention may be shaped into any number ofdesirable configurations to satisfy any number of overall system,geometry or space restrictions. The matrices may be three-dimensionalmatrices shaped to conform to the dimensions and shapes of a laminarilyorganized luminal organ or tissue structure. For example, in the use ofthe polymeric matrix for bladder reconstruction, a three-dimensionalmatrix may be used that has been shaped to conform to the dimensions andshapes of the whole or a part of a bladder. Naturally, the polymericmatrix may be shaped in different sizes and shapes to conform to thebladders of differently sized patients. Optionally, the polymeric matrixshould be shaped such that after its biodegradation, the resultingreconstructed bladder may be collapsible when empty in a fashion similarto a natural bladder. The polymeric matrix may also be shaped in otherfashions to accommodate the special needs of the patient. For example, apreviously injured or disabled patient, may have a different abdominalcavity and may require a bladder replacement scaffold, a bladderaugmentation scaffold, a bladder conduit scaffold, and a detrusor muscleequivalent scaffold adapted to fit.

In one aspect, the present invention contemplates additional scaffoldssuitable for use with the smooth muscle cell populations describedherein. For example, scaffolds suitable for implantation into the lungmay be provided.

A. Augmentation or Replacement Scaffolds

In one other aspect, the polymeric matrix or scaffold is shaped toconform to part of a bladder. In one embodiment, the shaped matrix isconformed to replace at least about 50%, at least about 60%, at leastabout 70%, at least about 80%, at least about 90%, or at least about 95%of the existing bladder of a recipient. In one other aspect, thepolymeric matrix or scaffold is shaped to conform to 100% or all of abladder.

In one embodiment, the polymeric matrix comprises a first implantable,biocompatible, synthetic or natural polymeric matrix or scaffold havingat least two separate surfaces, and a second implantable, biocompatible,synthetic or natural polymeric matrix or scaffold having at least twoseparate surfaces, which are adapted to mate to each other and shaped toconform to at least a part of the luminal organ or tissue structure inneed of the treatment when mated. The first and second polymericmatrices may be formed from one integral unit subdivided into two ormore distinct parts, or from two or more distinct parts, adapted tomate. In some embodiments, the first and second polymeric matrices oncemated may be used for reconstruction, repair, augmentation, orreplacement of a luminal organ or tissue structure.

In some embodiments, the first and second polymeric matrices aresymmetrical, while in other embodiments, the first and second polymericmatrices are asymmetrical. In one embodiment, the first polymeric matrixor scaffold has a hemispherical or quasi-hemispherical shape having aclosed, domed end and an open, equatorial border, and the secondpolymeric matrix or scaffold is a collar adapted to mate with theequatorial border of the first polymeric matrix. In another embodiment,the first and second polymeric matrices are each hemispherical orquasi-hemispherical in shape, having a closed, domed end and an open,equatorial border. In yet another embodiment, the first and secondpolymeric matrices each comprise a circular or semi-circular base and atleast 2 petals radially extending from each base. In this embodiment,the bases and petal shaped portions of the first and the secondpolymeric matrices are mated to create a hollow spherical orquasi-spherical matrix or scaffold such that a flanged longitudinal,elliptical opening is created on one side of the mated polymericmatrices, and a circular opening is created on the side opposite thelongitudinal opening. In another embodiment, the first and secondpolymeric matrices are made from 3 parts comprising a top, a front and asidepiece, adapted to mate. In this embodiment, the 3 distinct parts aremated using at least 3, preferably four vertical seams, thereby forminga crown shaped neo-bladder construct. The crown shaped constructs arepreferably used alone as a device for luminal organ reconstruction,repair, augmentation, or replacement. In one embodiment, the constructis a bladder augmentation scaffold. One example of a bladderaugmentation scaffold is depicted in FIG. 1. In another embodiment, theconstruct is a bladder replacement scaffold. One example of a bladderreplacement scaffold is depicted in FIG. 2.

Additionally, the first polymeric matrix, the second polymeric matrix,or both, may contain at least one receptacle or port adapted to receivea tubular vessel or insert where the connection of the construct to anative vessel or tube is necessary. The vessels or inserts arethemselves, for example, cylindrical or tubular shaped polymer matrices,each having at least one flange located at a first end of thecylindrical polymer. The vessels or inserts are, preferably, composed ofthe same biocompatible material as the first or second polymericmatrices described above. In some embodiments, the vessel or insert alsocontains a washer adapted to fit around the cylindrical or tubularvessel or insert polymer matrix. For example, the washer is a hydrogel.The cylindrical or tubular vessel or insert may optionally contain awasher. The washer may be hydrogel. Additionally, the cylindrical ortubular insert may be self-stabilizing.

In another embodiment, the receptacles or ports adapted to receivetubular vessels or inserts where the connection of the scaffold ormatrix (once seeded with cells) to a native vessel or tube is necessaryalso applies to other the matrices discussed below.

B. Urinary Diversions

The present invention provides neo-urinary diversion or conduitscaffolds that can be seeded with cells and used as a replacement forgastrointestinal tissue in the construction of a urinary diversion in asubject. For example, the neo-urinary diversions described herein mayhave application after radical cystectomy for the treatment of patientswho would otherwise undergo an ileal loop diversion.

In one aspect, the present invention contemplates conduit scaffolds ormatrices suitable for use as urinary diversions in a subject in needformed from the methods described herein. One end of the conduitscaffold may be connected to one or more ureters and the other end maybe connected to a urine reservoir that is external to the subject'sbody. In one embodiment, the conduit may exit the subject's body via astoma. In another embodiment, the polymeric matrix comprises a firstimplantable, biocompatible, synthetic polymeric matrix or scaffoldprovided in a tubular form. In some embodiments, the tubular scaffoldcomprises a first end configured to connect to a ureter of the subject.In another embodiment, the first scaffold further includes a second endconfigured to form a stoma or sphincter in the subject. In anotherembodiment, the first scaffold further includes at least one sideopening configured to connect to a least one ureter. In someembodiments, the first scaffold includes a first side opening configuredto attach to a first ureter and a second side opening configured toattach to a second ureter.

In one other embodiment, the tubular structure comprises a first endcomprising an even edge and a second end comprising a non-uniform oruneven edge. The non-uniform edge may include a circular base with anumber of petals radially extending from the base. The number of petalsmay be 1, 2, 3, 4, 5, or 6. The uneven edge may comprise a series ofpetals such as, for example, those shown in FIG. 3. In one embodiment,the tubular structure has a form suitable for use as a urinary diversionsystem or a conduit in a patient in need. In another embodiment, thesystem diverts urine from one or more ureters to an abdominal wallsection such as, for example, in the case of a ureterostomy. In otherembodiments, the system diverts urine from the bladder to an abdominalwall section such as, for example, in the case of a cystostomy. In oneother embodiment, the system connects the bladder to the urethra. In yetanother embodiment, a first system may divert urine from one or moreureters to an abdominal wall section and a second system may diverturine from the bladder to an abdominal wall section. In all embodiments,the system may divert urine from one or more ureters to an abdominalwall section such as, for example, in the formation of a stoma.

In another embodiment, the tubular matrix or scaffold is a urinarydiversion or conduit scaffold.

In one embodiment, the tubular structure of the urinary diversion systemis of rectangular, circular, or triangular cross sectional area. FIG. 3Aillustrates some of the different cross sectional configurationscontemplated herein.

In another embodiment, tubular structure retains sufficient rigidity toremain patent following implantation. In one other embodiment, thetubular structure's rigidity is retained with or without the use of acatheter in its lumen. Where a catheter is used, it can be placed intothe luminal space of the tubular structure to provide additionalpatency.

In one other embodiment, the conduit scaffold may further include asecond scaffold in the form of a round or ovoid connector configured toconnect the first end of the first scaffold to a ureter. In yet anotherembodiment, the conduit scaffold may further include a third scaffold inthe form of a washer-ring configured to form a stoma or sphincter withthe second end of the first tubular scaffold to create a stoma in asubject. FIG. 3B illustrates variations of a urinary diversion construct(A—open claim ovoid; B—open claim ovoid receptacle; C—closed ovoidreceptacle and three tubes).

In some embodiments, the tubular structure may include a washerstructure for connection to a tissue, organ or body part to achieveanastomosis for the creation of a continent stoma or sphincter. Inanother embodiment, the washer is provided with a thickness of aboutless than 1 mm, about less than 1.5 mm, about less than 2 mm, about lessthan 2.5 mm, about less than 3 mm, about less than 3.5 mm, about lessthan 4 mm, about less than 4.5 mm, or about less than 5 mm.

In one embodiment, the urinary diversion or conduit scaffold is shapedinto the configuration shown in FIG. 3.

In one other embodiment, the tubular structure comprises a first endcomprising an even edge and a second end comprising a non-uniform oruneven edge. The non-uniform edge may include one or more fastenersconfigured for attachment to an external region of the subject, such asin the formation of a stoma external to the subject. In one embodiment,the first and second ends of the tubular structure may be in the formillustrated in FIG. 3. The number of fasteners may be 1, 2, 3, 4, 5, or6.

In another embodiment, the tubular scaffold is in the form depicted inFIG. 27.

FIG. 4A depicts a part of the normal anatomy for the human urinarysystem.

In one embodiment, the tubular structure has a form suitable for use asa urinary diversion or a conduit in a patient in need. In anotherembodiment, the conduit diverts urine from one or more ureters to anabdominal wall section such as, for example, in the case of aureterostomy (FIG. 4D). In other embodiments, the conduit diverts urinefrom the bladder to an abdominal wall section such as, for example, inthe case of a cystostomy (FIG. 4B). In one other embodiment, the conduitconnects the bladder to the urethra (FIG. 4D). In yet anotherembodiment, a first conduit may divert urine from one or more ureters toan abdominal wall section and a second conduit may divert urine from thebladder to an abdominal wall section. In all embodiments, the conduitmay divert urine from one or more ureters to an abdominal wall section(FIG. 4B). In all embodiments, the conduit may be configured to form astoma.

In one embodiment, the tubular structure of the urinary diversion orconduit scaffold is of rectangular, circular, or triangular crosssectional area. In another embodiment, the tubular structure retainssufficient rigidity to remain patent following implantation. In oneother embodiment, the tubular structure's rigidity is retained with orwithout the use of a catheter in its lumen. In some embodiments, aurinary diversion scaffolds further include a catheter configured to beplaced in the luminal space of tubular structure upon implantation. Inone embodiment, the catheter is a Foley-like balloon catheter. Where acatheter is used, it can be placed into the luminal space of the tubularstructure to provide additional patency. Those of ordinary skill in theart will appreciate that other catheters known in the art may besuitable for use with the present invention.

In another embodiment, the thickness of the tubular wall of thescaffolds will be less than about 2 mm, less than about 2.5 mm, lessthan about 3.5 mm, less than about 4 mm, less than about 4.5 mm, lessthan about 5 mm, less than about 5.5 mm, or less than about 6 mm.

In some embodiments, the scaffolds may have variable outer and innerdiameters. In one embodiment, the ends of the scaffold may be flared,non-flared, sealed, or rounded.

In other embodiments, the scaffold is permeable to urine. In oneembodiment, the scaffold's pore size is about greater than about 0microns to about 500 microns. In another embodiment, the pore size isfrom about 100 microns to about 200 microns. In another embodiment, thepore size is from about 150 microns to about 200 microns. In otherembodiments, the pore size is about 100 microns, about 110 microns,about 120 microns, about 130 microns, about 140 microns, about 150microns, about 160 microns, about 170 microns, about 180 microns, about190 microns, or about 200 microns. In some embodiments, the pore size isabout 100 microns, about 200 microns, about 300 microns, about 400microns, about 500 microns, or about 600 microns. In other embodiments,the scaffold includes a pore architecture that is a single pore sizedistribution, multiple pore size distribution, or a pore gradientdistribution.

In another embodiment, the scaffold material is suturable and may formconnections with tissue that are resistant to leakage.

In other embodiments, the tubular scaffold material is selected tomaintain patency throughout the duration of implantation use, supportcell attachment and the in-growth of host tissue, and retainflexibility. In another embodiment, the material will have a burststrength that exceeds the pressures to which it will be exposed duringnormal in vivo fluid cycling. In other embodiments, the material willhave a degradation time commensurate with host tissue in-growth.

C. Muscle Equivalents

In one aspect, the polymeric matrix or scaffold of the present inventionis a muscle equivalent scaffold. In one embodiment, the muscleequivalent scaffold is a detrusor muscle equivalent scaffold. In anotherembodiment, the scaffold is suitable for laparoscopic implantation.

In one aspect, the polymeric matrix comprises a polymeric matrix orscaffold shaped to conform to at least a part of the organ or tissuestructure in need of said treatment and of a sufficient size to belaparoscopically implanted. In certain embodiments, the polymeric matrixor scaffold of the invention is between about 3 and about 20 cm inlength. In one embodiment the polymeric matrix or scaffold is about 20cm in maximal length. In another embodiment, the polymeric matrix orscaffold is about 15 cm in maximal length. In another embodiment, thepolymeric matrix or scaffold is about 10 cm in maximal length. Inanother embodiment, the polymeric matrix or scaffold is about 8 cm inmaximal length. In another embodiment, the polymeric matrix or scaffoldis about 4 cm in maximal length. In yet another embodiment, thepolymeric matrix or scaffold is about 3 cm in maximal length. In certainembodiments, the polymeric matrix or scaffold of the invention isbetween about 1 and about 8 cm in width. In some embodiments, thepolymeric matrix or scaffold is about 4 cm in maximal width. In otherembodiments, the polymeric matrix or scaffold is about 3 cm in maximalwidth. In yet other embodiments, the polymeric matrix or scaffold isabout 5 cm in maximal width.

In one embodiment, the polymeric matrix or scaffold has athree-dimensional (3-D) shape. In another embodiment, the polymericmatrix or scaffold has a flat shape. In one embodiment, the flat-shapedpolymeric matrix or scaffold comprises pre-treated areas to allow moreflexibility. In certain embodiments, the pre-treated areas are coated inthe areas to be creased. In one embodiment, the polymeric matrix orscaffold is sufficiently malleable to be rolled, folded, or otherwiseshaped for implantation through a laparoscope tube and/or port. In suchembodiments, the polymeric matrix or scaffold is sufficiently malleableto be unrolled, unfolded, or otherwise returned to shape followinginsertion through the laparoscope tube and/or port. In one embodiment,the polymeric matrix or scaffold is cut into 2, 3, 4, 5, 6, 7, 8, 9 or10 strips prior to implantation through a laparoscope tube and/or port.In certain embodiments, the 2, 3, 4, 5, 6, 7, 8, 9 or 10 strips aremated prior to implantation through a laparoscope tube and/or port. The2, 3, 4, 5, 6, 7, 8, 9 or 10 strips may be mated using glue, staples,sutures, or other technique known to one of ordinary skill in the art.In such embodiments the 2, 3, 4, 5, 6, 7, 8, 9 or 10 mated strips arefolded and/or stacked to pass through a laparoscope tube and/or port. Insuch embodiments, the 2, 3, 4, 5, 6, 7, 8, 9 or 10 strips are unfoldedand/or unstacked following insertion through the laparoscope tube and/orport. In some embodiments, the previously placed mating means aretightened as appropriate following insertion through the laparoscopetube and/or port.

In one embodiment, the polymeric matrix comprises a first implantable,biocompatible, synthetic or natural polymeric matrix or scaffoldprovided in the form of a patch or in the form of a strip. In oneembodiment, the patch has a form suitable for use as a detrusor muscleequivalent in the bladder of a patient in need. In one other embodiment,the patch has a form suitable for increasing the volume capacity of theexisting bladder of a patient in need. In certain embodiments, the patchincreases the bladder size between about 50 mL and about 500 mL. In someembodiments, the patch would increase bladder size in increments of 50mL. In some embodiments, the patch increases the bladder size about 450mL. In one embodiment, a surface area increase of 30 cm² increases thevolume of a 200 mL bladder to 250 mL. In another embodiment, an increaseof 25 cm² increases the volume of a 350 mL bladder to 400 mL. In oneembodiment, the scaffold has a two-dimensional surface area of about 30cm². In another embodiment, the scaffold has a two-dimensional surfacearea of about 25 cm². In one embodiment, the patch is in the form of astrip, disc, square, ellipsoid, or any other appropriate configuration.In other embodiments, the patch is provide in a pre-folded form, e.g.,like an accordion.

FIG. 5A-B show examples of a muscle equivalent scaffold or polymericmatrix. In one embodiment, the polymeric matrix or scaffold is in theshape of a double wedge, e.g., the shape shown in FIG. 5A. In anotherembodiment, the polymeric matrix is shaped into one of theconfigurations shown in FIGS. 6-9.

In all embodiments, the polymeric matrix or scaffold is shaped so as tominimize the strain on both the bladder and matrix or scaffold.

In another embodiment, the polymeric matrix comprises a firstimplantable, biocompatible, synthetic or natural polymeric matrix orscaffold provided in the form of a patch or in the form of a strip. Inone embodiment, the patch has a form suitable for use as a detrusormuscle equivalent in the bladder of a patient in need. In one otherembodiment, the patch has a form suitable for increasing the volumecapacity of the existing bladder of a patient in need. In someembodiments, the patch would increase bladder size in increments of 50mL. In one embodiment, the patch is in the form of a strip, disc,square, ellipsoid, or any other appropriate configuration. In otherembodiments, the patch is provide in a pre-folded form, e.g., like anaccordion.

In one embodiment, the polymeric matrix is shaped into one of theconfigurations shown in FIG. 1-9 or 27.

In all embodiments, the biocompatible material used for these matricesor scaffolds is, for example, biodegradable. In all the embodiments, thebiocompatible material may be polyglycolic acid.

In all embodiments, the polymeric matrix or scaffold is coated with abiocompatible and biodegradable shaped setting material. In oneembodiment, the shape setting material may comprise a liquid copolymer.In another embodiment, the liquid co-polymer may comprise a liquefiedlactide/glycolide copolymer. In one embodiment, the liquid co-polymermay comprise poly-DL-lactide-co-glycolide.

5. Constructs

In one aspect, the invention provides one or more polymeric scaffolds ormatrices that are seeded with at least one cell population. Suchscaffolds that have been seeded with a cell population and may bereferred to herein as “constructs”. In one embodiment, the cell-seededpolymeric matrix or matrices form a neo-bladder construct selected fromthe group consisting of a bladder replacement construct, a bladderaugmentation construct, a bladder conduit construct, and a detrusormuscle equivalent construct.

Those of skill in the art will appreciate that the seeding or depositionof one or more cell populations described herein may be achieved byvarious methods known in the art. For example, bioreactor incubation andculturing, (Bertram et al. U.S. Published Application 20070276507;McAllister et al. U.S. Pat. No. 7,112,218; Auger et al. U.S. Pat. No.5,618,718; Niklason et al. U.S. Pat. No. 6,537,567); pressure-inducedseeding (Torigoe et al. (2007) Cell Transplant., 16(7):729-39; Wang etal. (2006) Biomaterials. May; 27(13):2738-46); and electrostatic seeding(Bowlin et al. U.S. Pat. No. 5,723,324) may be used. In addition, arecent technique that simultaneously coats electrospun fibers with anaerosol of cells may be suitable for seeding or deposition (Stankus etal. (2007) Biomaterials, 28:2738-2746).

In one embodiment, the deposition of cells includes the step ofcontacting a scaffold with a cell attachment enhancing protein. Inanother embodiment, the enhancing protein is one or more of thefollowing: fibronection, collagen, and MATRIGEL™. In one otherembodiment, the scaffold is free of a cell attachment enhancing protein.In another embodiment, the deposition of cells includes the step ofculturing after contacting a scaffold with a cell population. In yetanother embodiment, the culturing may include conditioning by pulsatileand/or steady flow in a bioreactor.

Smooth muscle cell populations isolated from adipose or peripheral bloodas described herein may then be seeded on a scaffold described herein.

The following is a representative example of a protocol for seedingcells on a scaffold. Adipose- or peripheral blood-derived smooth musclecells may be expanded for up to 7 weeks to generate the quantity ofcells required for seeding a scaffold. The density of cells suitable forseeding a scaffold is described below. Adipose-derived smooth musclecells may be expanded for 2 passages before harvesting of cells forseeding of scaffolds to produce a construct. Peripheral blood-derivedsmooth muscle cell cultures may be expanded to P3-4 before harvestingfor scaffold seeding. To prepare a scaffold for cell seeding, a suitablematerial (e.g., PGA felt) may be cut to size, sutured into theappropriate shape, and coated with material (e.g., PLGA). The scaffoldmay then be sterilized using a suitable method (e.g., ethylene oxide).On the day prior to cell seeding, the sterilized scaffold may beserially pre-wetted by saturation with 60% ethanol/40% D-PBS, 100%D-PBS, D-MEM/10% FBS or α-MEM/10% FBS followed by incubation inD-MEM/10% FBS or α-MEM/10% FBS at room temperature overnight. Thescaffold can then be seeded with adipose-, or peripheral blood-derivedsmooth muscle cells and the seeded construct matured in a humidified 37°C. incubator at 5% CO₂ until implantation in a subject (e.g., by day 7).Those of ordinary skill in the art will appreciate additional methodsfor preparing scaffolds for seeding of cells and seeding of cells ontoscaffolds.

In one aspect, the present invention provides methods of preparing aconstruct in a reduced time frame, which is advantageous to the subjectawaiting implantation of a construct. It has been reported thatundifferentiated adipose stem cells derived from SVF must be incubatedin inductive media for 6 weeks prior to differentiation into smoothmuscle cells (Jack et al. 2009 supra). In one embodiment, the methodincludes the steps of a) obtaining a human adipose tissue sample; b)isolating a fully differentiated smooth muscle cell population from thesample; c) culturing the cell population; and d) contacting the cellpopulation with a shaped polymeric matrix cell construct, wherein stepsa), b), c) and d) are performed in about 45 days or less. In anotherembodiment, the isolating step is performed without cell selection. Inanother embodiment, the isolating step b) is performed about 72 hours orless after obtaining step a). In yet another embodiment, the culturingstep c) is performed in about 4 weeks or less. In other embodiments, thecontacting step d) is performed in about 10 days or less. In anotherembodiment, steps a), b), c) and d) are performed in about 28 days orless. In one other embodiment, the isolating step b) is performed about48 hours or less after obtaining step a). In one embodiment, theculturing step c) is performed in about 2 weeks or less. In anotherembodiment, the contacting step d) is performed in about 5 days or less.In all embodiments, the human adipose tissue sample is obtained from anautologous source. In one other embodiment, the method further includesthe step of detecting expression of a smooth muscle cell marker. Inanother embodiment, expression is mRNA expression. In a furtherembodiment, the expression is polypeptide expression. In one embodiment,the polypeptide expression is detected by intracellularimmunoflourescence.

In one embodiment, the scaffold comprises a cell population as describedherein. In another embodiment, the scaffold consists essentially of acell population as described herein. In one other embodiment, thescaffold consists of a cell population as described herein.

The first polymeric matrix or the second polymeric matrix, if any, orboth, comprise at least one cell population deposited on or in a firstsurface of the first polymeric matrix, a first surface of the secondpolymeric matrix, or both, to form a construct of matrix or scaffoldplus cells, wherein at least one cell population comprises substantiallya muscle cell population. The muscle cell population is, e.g., a smoothmuscle cell population. In a preferred embodiment, the first surface andthe second surface are each the outer surface of the first and secondpolymeric matrices.

In another embodiment, the construct containing the matrix and cells isfree of any other cell populations. In a preferred embodiment, theconstruct is free of urothelial cells.

These constructs are used to provide a luminal organ or tissuestructures such as genitourinary organs, including for example, theurinary bladder, ureters and urethra, to a subject in need. The subjectmay require the reconstruction, repair, augmentation or replacement ofsuch organs or tissues. In one embodiment, the luminal organ or tissuestructure is a bladder or portion thereof, and the polymeric matrix orscaffold has smooth muscle cells deposited on a surface of the matrix.The constructs may also be used to provide a urinary diversion orconduit, or a detrusor muscle equivalent.

In one aspect, the invention provides urinary diversion or conduitscaffolds or matrices that are seeded with a cell population describedherein. Such scaffolds that have been seeded with a cell population andmay be referred to herein as “constructs”. In one embodiment, theurinary diversion or bladder conduit construct is made up of one or morescaffolds as described herein and a cell population deposited on one ormore surfaces of the one or more scaffolds as described herein.

In one aspect, the present invention provides muscle equivalentconstructs that may be used to enhance an existing luminal organ ortissue structures such as genitourinary organs, including for example,the urinary bladder, to a subject in need. The subject may requireexpansion or treatment of such organs or tissues. In one embodiment, theluminal organ or tissue structure is a bladder or portion thereof, andthe polymeric matrix or scaffold has smooth muscle cells deposited on asurface of the matrix. In one embodiment, the constructs are used toprovide a detrusor muscle equivalent.

Those of ordinary skill in the art will appreciate there are severalsuitable methods for depositing cell populations upon matrices orscaffolds.

In one aspect, the constructs are suitable for implantation into asubject in need of a new organ or tissue structure. In one embodiment,the construct comprises a population of cells that produce the cytokineMCP-1. In another embodiment, the MCP-1 elicits the migration of thesubject's or recipient's native mesenchymal stem cells to the site ofimplantation. In one embodiment, the migrating recipient nativemesenchymal stem cells assist in the regeneration of the new organ ortissue structure.

In one other aspect, the invention provides scaffolds seeded with cellsat particular cell densities. In one embodiment, a scaffold is seededwith a smooth muscle cell population at a cell density of about 20×10⁶to about 30×10⁶ cells. In another embodiment, the cell density is about1×10⁶ to about 40×10⁶, about 1×10⁶ to about 30×10⁶, about 1×10⁶ to about20×10⁶, about 1×10⁶ to about 10×10⁶, or about 1×10⁶ to about 5×10⁶.

In a further embodiment, the density is about 20×10⁶ to about 98×10⁶cells. In yet further embodiments, the density is about 21×10⁶ to about97×10⁶, about 22×10⁶ to about 95×10⁶, about 23×10⁶ to about 93×10⁶,about 24×10⁶ to about 91×10⁶, about 25×10⁶ to about 89×10⁶, about 26×10⁶to about 87×10⁶, about 28×10⁶ to about 85×10⁶, about 29×10⁶ to about83×10⁶, about 30×10⁶ to about 80×10⁶, about 35×10⁶ to about 75×10⁶,about 40×10⁶ to about 70×10⁶, about 45×10⁶ to about 65×10⁶, or about50×10⁶ to about 60×10⁶. In a preferred embodiment, the density is about24×10⁶ to about 91×10⁶ cells

In another embodiment, the density is about 2.5×10⁶ to about 40×10⁶,about 5×10⁶ to about 40×10⁶, about 7.5×10⁶ to about 35×10⁶, about 10×10⁶to about 30×10⁶, about 15×10⁶ to about 25×10⁶, and about 17.5×10⁶ toabout 22.5×10⁶. In another embodiment, the cell density is about 1×10⁶,about 2×10⁶, about 3×10⁶, about 4×10⁶, about 5×10⁶, about 6×10⁶, about7×10⁶, about 8×10⁶, about 9×10⁶, about 10×10⁶, about 11×10⁶, about12×10⁶, about 13×10⁶, about 14×10⁶, about 15×10⁶, about 16×10⁶, about17×10⁶, about 18×10⁶, about 19×10⁶, about 20×10⁶, about 21×10⁶, about22×10⁶, about 23×10⁶, about 24×10⁶, about 25×10⁶, about 26×10⁶, about27×10⁶, about 28×10⁶, about 29×10⁶, about 30×10⁶, about 31×10⁶, about32×10⁶, about 33×10⁶, about 34×10⁶, about 35×10⁶, about 36×10⁶, about37×10⁶, about 38×10⁶, about 39×10⁶, about 40×10⁶, about 41×10⁶, about42×10⁶, about 43×10⁶, about 44×10⁶, about 45×10⁶, about 46×10⁶, about47×10⁶, about 48×10⁶, about 49×10⁶, about 50×10⁶, about 51×10⁶, about52×10⁶, about 53×10⁶, about 54×10⁶, about 55×10⁶, about 56×10⁶, about57×10⁶, about 58×10⁶, about 59×10⁶, about 60×10⁶, about 61×10⁶, about62×10⁶, about 63×10⁶, about 64×10⁶, about 65×10⁶, about 66×10⁶, about67×10⁶, about 68×10⁶, about 69×10⁶, about 70×10⁶, about 71×10⁶, about72×10⁶, about 73×10⁶, about 74×10⁶, about 75×10⁶, about 76×10⁶, about77×10⁶, about 78×10⁶, about 79×10⁶, about 80×10⁶, about 81×10⁶, about82×10⁶, about 83×10⁶, about 84×10⁶, about 85×10⁶, about 86×10⁶, about87×10⁶, about 88×10⁶, about 89×10⁶, about 90×10⁶, about 91×10⁶, about92×10⁶, about 93×10⁶, about 94×10⁶, about 95×10⁶, about 96×10⁶, about97×10⁶, about 98×10⁶, or about 99×10⁶.

In a further aspect, the invention provides scaffolds seeded with cellsat particular cell densities per cm² of a scaffold. In one embodiment,the density is about 3,000 cells/cm² to about 15,000 cells/cm², about3,500 cells/cm² to about 14,500 cells/cm², about 4,000 cells/cm² toabout 14,000 cells/cm², about 4,500 cells/cm² to about 13,500 cells/cm²,about 5,000 cells/cm² to about 13,000 cells/cm², about 4,500 cells/cm²to about 13,500 cells/cm², about 5,000 cells/cm² to about 13,000cells/cm², about 5,500 cells/cm² to about 12,500 cells/cm², about 6,000cells/cm² to about 12,000 cells/cm², about 6,500 cells/cm² to about11,500 cells/cm², about 7,000 cells/cm² to about 11,000 cells/cm², about7,500 cells/cm² to about 10,500 cells/cm², about 8,000 cells/cm² toabout 10,000 cells/cm², about 7,500 cells/cm² to about 9,500 cells/cm²,or about 8,000 cells/cm² to about 9,000 cells/cm². In a preferredembodiment, the density is about 3,000 cells/cm² to about 7,000cells/cm², or about 9,000 cells/cm² to about 15,000 cells/cm².

In one aspect, the constructs of the present invention are adapted toprovide particular features to the subject following implantation. Inone embodiment, the constructs are adapted to provide regeneration tothe subject following implantation. In another embodiment, theconstructs are adapted to promote regeneration in a subject at the siteof implantation. For example, following implantation, regenerated tissuemay form from the construct itself at the site of implantation. Inanother embodiment, the construct may impart functional attributes tothe subject following implantation. For example, a urinary diversionconstruct may be adapted to allow the passage of a subject's urine froma first ureter (e.g., first side opening) to the interior of the tubularscaffold, and/or adapted to provide temporary storage and passage ofurine (e.g., tubular scaffold) out of a subject. In one embodiment, aurinary diversion construct may be adapted to provide an epithelializedmucosa upon implantation. In another embodiment, a construct may beadapted to provide homeostatic regulative development of a new organ ortissue structure in a subject.

6. Methods of Use

In one aspect, the present invention contemplates methods for providinga laminarily organized luminal organ or tissue structure to a subject inneed of such treatment. In one embodiment, the subject may be in need ofreconstruction, repair, augmentation, or replacement of an organ ortissue. In one embodiment, the method includes the step of providing abiocompatible synthetic or natural polymeric matrix shaped to conform toat least a part of the organ or tissue structure in need of an organ ortissue structure. The providing step may be followed by depositing atleast one cell population that is not derived from the organ or tissuestructure that is the subject of the reconstruction, repair,augmentation or replacement. The depositing step may include culturingthe cell population on the polymeric matrix. After depositing the cellpopulation on the matrix to provide a construct, it can be implantedinto a patient at the site of treatment for the formation of the desiredlaminarily organized luminal organ or tissue structure. In oneembodiment, the laminarly organized luminal organ or tissue structure isa bladder or a part of a bladder.

In one other aspect, the present invention provides methods forproviding a laminarily organized luminal organ or tissue structure to asubject in need. In one embodiment, the method includes the steps of a)providing a biocompatible synthetic or natural polymeric matrix shapedto conform to at least a part of the organ or tissue structure in needof said treatment; b) depositing on or in a first area of the polymericmatrix an autologous cell population that is not derived from a nativeorgan or tissue corresponding to the new organ or tissue structure; andc) implanting the shaped polymeric matrix cell construct into said thesubject for the formation of laminarily organized luminal organ ortissue structure. In one other aspect, the present invention providesmethods for providing a neo-bladder or portion thereof to a subject inneed. In one embodiment, the method includes a) providing abiocompatible synthetic or natural polymeric matrix shaped to conform toa bladder or portion thereof; b) depositing an autologous cellpopulation that is not derived from the subject's bladder on or in afirst area of the polymeric matrix; and c) implanting the shapedpolymeric matrix cell construct into the subject for the formation ofthe neo-bladder or portion thereof. In another embodiment, the cellpopulation of step b) of the methods described herein contains one ormore peripheral blood-derived smooth muscle cells having contractilefunction that are positive for a smooth muscle cell marker, or the cellpopulation of step b) contains one or more adipose tissue-derived smoothmuscle cells having contractile function that are positive for a smoothmuscle cell marker. In one other embodiment, the contractile function ofthe cell population is calcium-dependent.

In one embodiment, the methods of the present invention further includethe step of wrapping the implanted conduit construct with the subject'somentum, mesentery, muscle fascia, and/or peritoneum to allow forvascularization.

In one other aspect, the present invention provides methods forproviding a urinary diversion or conduit for a defective bladder in asubject in need. In one embodiment, the method for providing a urinarydiversion to a subject in need includes the steps of (a) providing abiocompatible conduit scaffold; (b) depositing a first cell populationon or in a first area of said scaffold, said first cell population beingsubstantially a muscle cell population; and (c) implanting the scaffoldof step (b) into said subject to form a conduit that allows urine toexit the subject. In another embodiment, the biocompatible material isbiodegradeable. In other embodiments, the biocompatible material ispolyglycolic acid. In yet another embodiment, the first cell populationis substantially a smooth muscle cell population.

In one embodiment, the method includes the step of providing a urinarydiversion or conduit scaffold as described herein. In other additionalembodiments, the urinary diversion or conduit scaffold is provided inmultiple parts, such as a first, second, and third scaffold, asdescribed herein. In another embodiment, the method further includes thestep of depositing a cell population that is not derived from thedefective bladder to form a urinary diversion or conduit construct. Inone other embodiment, the depositing step may include culturing the cellpopulation on the scaffold. In some embodiments, the methods furtherincludes the step of implanting the urinary diversion construct into apatient in need. In another embodiment, the implantation is at the siteof the defective bladder.

In one embodiment, an open end of the construct (e.g., a first endconfigured to connect to the abdominal wall) is anastomosed to the skin(ostomy) through the abdominal or suprapubic wall to form a stoma orsphincter. In another embodiment, a catheter is inserted through stomaopening and into the lumen of the construct to provide urine outflow.

FIG. 10 illustrates a configuration for an implanted conduit construct.

In another embodiment, the methods of the present invention furtherinclude the step of monitoring the conduit for the presence of anobstruction following implantation of the urinary diversion construct.The obstruction may be caused by the build-up of detritis. The methodmay further include the step of removing detritis from the lumen of theconduit if an obstruction is detected.

In one aspect, the present invention provides a urinary diversion to asubject in need on a temporary basis. In one embodiment, a temporaryurinary diversion or conduit construct is implanted into a subject toform a stoma opening, and a catheter or other device is temporarilyinserted through the stoma to the lumen of the conduit construct. Atemporary conduit provides the advantage of allowing urine to exit thesubject while a permanent solution to the defective bladder isattempted. For example, the implantation of a conduit construct could beperformed prior to, following, or simultaneous with the implantation ofa neo-bladder construct seeded with a cell population (see for exampleBertram et al. supra). FIG. 11 shows an example of the implantedcomponents of a temporary urinary diversion construct.

In one embodiment, the methods of the present invention further includethe step of wrapping the implanted urinary diversion or conduitconstruct with the subject's omentum, mesentery, muscle fascia, and/orperitoneum to allow for vascularization.

In one aspect, the present invention provides a urinary diversion to asubject in need on a permanent basis. FIG. 12 shows an example of theimplanted components of a permanent urinary diversion construct.

In one embodiment, the constructs described herein may be used for aprostatic urethra replacement and urinary diversion. Such a procedure isnecessary for subjects requiring a radical prostatectomy to remove theprostatic urethra. In other embodiments, the constructs may be used fora percutaneous diversion tube to form a continent tube with a valve-likekink In an additional embodiment, the constructs may be used as abladder neck sling and wrapping materials used in bladder neck surgeryand urinary outlets with continent channels or catherizable openings.Examples of such embodiments are depicted in FIG. 13.

In one aspect, the urinary diversion constructs of the present inventionprovide an epithelialized mucosa. In one embodiment, the construct isadapted to form an epithelialized mucosa upon implantation. In oneembodiment, the epithelialized mucosa comprises a vestibular region anda mucocutaneous region. In another embodiment, the vestibular region isadjacent to the mucocutaneous region. In another embodiment, themucocutaneous region is located at the stromal end of the constructconnected to the abdominal wall and skin of the subject. In general,naturally-occurring mucocutaneous regions are characterized by thepresence of mucosa and cutaneous skin and typically exist near theorifices of the body where the external skin ends and the mucosa thatcovers the inside of the body starts. The epithelialized mucosa providedby the constructs and methods of the present invention develops at thefirst end of the urinary diversion construct following implantation intothe subject. In a further embodiment, the epithelialized mucosa ischaracterized by the presence of an epithelium that first appears in thevestibular region and gradually expands or increases through themucocutaneous region towards the stomal end of the construct. In anotherembodiment, the epithelium is characterized by expression of anepithelial cell marker. In a further embodiment, the epithelial cellmarker is cytokeratin. The cytokeratin may be one or more of thecytokeratins known in the art including, without limitation,cytokeratins 1 through 19. In one other embodiment, the cytokeratin isdetectable with AE-1/AE3 antibody.

Grafting of scaffolds to an organ or tissue to be enlarged can beperformed according to the methods described in the Examples oraccording to art-recognized methods. The matrix or scaffold can begrafted to an organ or tissue of the subject by suturing the graftmaterial to the target organ.

The described techniques may be used to expand an existing laminarilyorganized luminal organ or tissue structure in a patient in need of suchtreatment. For example, an existing laminarily organized luminal organor tissue structure may be enlarged by providing a polymeric matrix orscaffold shaped to conform to at least a part of the organ or tissuestructure in need of said treatment and of a sufficient size to belaparoscopically implanted, depositing an autologous cell populationthat is not derived from the organ or tissue structure on or in a firstarea of said polymeric matrix; and laparoscopically implanting theshaped polymeric matrix construct into said patient at the site of saidtreatment such that the existing laminarily organized luminal organ ortissue structure is expanded.

FIG. 7 e depicts possible surgical methods for the implantation of amuscle equivalent scaffold described herein. FIG. 7 f depictsimplantation sites on an empty and full bladder. FIG. 7 g depicts aurinary bladder model with surgical slit showing ellipsoid created uponsectioning of surface. A plastic tube may be used as a model of thelimited space available in order to pass the folded or rolled polymericmatrices or scaffolds of the invention.

The described techniques may also be used to increase bladder volumetriccapacity in a patient in need of such treatment. For example, bladdervolumetric capacity may be increased by providing a biocompatiblesynthetic or natural polymeric matrix shaped to conform to at least apart of the organ or tissue structure in need of said treatment and of asufficient size to be laparoscopically implanted; depositing anautologous cell population that is not derived from the organ or tissuestructure on or in a first area of said polymeric matrix; andlaparoscopically implanting the shaped polymeric matrix constructlaparoscopically into said patient at the site of said treatment suchthat bladder volume capacity is increased. In one embodiment, the matrixor scaffold of the instant invention is suitable for increasing bladdervolume capacity about 50 mL. In other embodiments, the matrix orscaffold of the instant invention is suitable for increasing bladdervolume capacity about 100 mL. In other embodiments, the matrix orscaffold of the instant invention is suitable for increasing bladdervolume capacity about 60, about 70, about 80, or about 90 mL.

The described techniques may further be used to expand a bladderincision site in a patient in need of such treatment. For example, abladder incision site may be expanded by providing a biocompatiblesynthetic or natural polymeric matrix shaped to conform to at least apart of the organ or tissue structure in need of said treatment and of asufficient size to be laparoscopically implanted; b) depositing anautologous cell population that is not derived from the organ or tissuestructure on or in a first area of said polymeric matrix; and c)laparoscopically implanting the shaped polymeric matrix constructlaparoscopically into said patient at the site of said treatment suchthat the bladder incision site is expanded.

Another non-limiting use of the invention includes methods for thetreatment of urinary incontinence in a patient in need of suchtreatment. For example, urinary incontinence may be treated by providinga biocompatible synthetic or natural polymeric matrix shaped to conformto at least a part of the organ or tissue structure in need of saidtreatment and of a sufficient size to be laparoscopically implanted;depositing an autologous cell population that is not derived from theorgan or tissue structure on or in a first area of said polymericmatrix; and laparoscopically implanting the shaped polymeric matrixconstruct laparoscopically into said patient at the site of saidtreatment such that bladder volume capacity is increased.

In one embodiment, the scaffolds, cell populations, and methodsdescribed herein may further be used for the preparation of a medicamentuseful in the treatment of a disorder described herein. The disordersinclude any condition in a subject that requires the regeneration,reconstruction, repair, augmentation or replacement of laminarlyorganized luminal organs or tissue structures. In another embodiment,the organ or tissue structure is a bladder or a part of the bladder.

In another embodiment, the cells deposited on the implanted constructproduce MCP-1 and release it at the site of implantation, whichstimulates native mesenchymal stem cells (MSCs) to migrate to the siteof implantation. In one other embodiment, the native MSCs facilitateand/or enhance regeneration of the implanted construct at the site ofimplantation.

In one embodiment, the cell population deposited is a smooth muscle cell(SMC) population derived from peripheral blood or from adipose tissue asdescribed herein. In another embodiment, the SMC population includes atleast one cell that has contractile function and is positive for asmooth muscle cell marker, such as myocardin, alpha-smooth muscle actin,calponin, myosin heavy chain, BAALC, desmin, myofibroblast antigen,SM22, and any combination thereof. In other embodiments, the SMCpopulation includes at least one cell that demonstrates myocardin(MYOCD) expression. The MYOCD expression may be expression of a nucleicacid encoding a MYCOD polypeptide or a MYOCD polypeptide. In anotherembodiment, the contractile function of the SMC is calcium-dependent. Inone embodiment, the laminarily organized luminal organ or tissuestructure that is the subject of reconstruction, repair, augmentation orreplacement is a bladder or a portion of a bladder. In anotherembodiment, the polymeric matrix is free of urothelial cells.

In all embodiments, the methods of the present invention utilize aconstruct for implantation that is based upon a bladder replacementscaffold, a bladder augmentation scaffold, a bladder conduit scaffold,or a detrusor muscle equivalent scaffold that has been seeded with acell population as described herein.

In another embodiment, the methods for the regeneration, reconstruction,repair, augmentation or replacement of laminarly organized luminalorgans or tissue structures described herein include the steps of a)providing a biocompatible synthetic or natural polymeric matrix shapedto conform to at least a part of the luminal organ or tissue structurein need of said treatment; b) depositing a first cell population on orin a first area of said polymeric matrix at a cell density describedherein, said first cell population being substantially a muscle cellpopulation; and c) implanting the shaped polymeric matrix cell constructinto said patient at the site of said treatment for the formation of thelaminarily organized luminal organ or tissue structure. In one otherembodiment, the laminarily organized luminal organ or tissue structureformed in vivo exhibits the compliance of natural bladder tissue.

In one other aspect, the present invention provides methods for theregeneration of a neo-bladder following implantation into a subject inneed thereof based upon biomechanical stimulation or cycling. In oneaspect, the methods are suitable for use in promoting the regenerationof an implanted neo-bladder construct that has been implanted for theaugmentation or replacement of a bladder or a portion of a bladder. Inone embodiment, the neo-bladder construct is formed from seeding cellson a neo-bladder matrix or scaffold. In another embodiment, theneo-bladder scaffold is a bladder replacement scaffold, a bladderaugmentation scaffold, a bladder conduit scaffold, or a detrusor muscleequivalent scaffold.

In one aspect, the method of the present invention applies to implantedneo-bladder constructs formed from seeding neo-bladder scaffolds with atleast one cell population. In one embodiment, the cell-seeded polymericmatrix (or matrices) is a bladder replacement scaffold, a bladderaugmentation scaffold, a bladder conduit scaffold, or a detrusor muscleequivalent scaffold. In one embodiment, the at least one cell populationcomprises substantially a muscle cell population. In another embodiment,the muscle cell population may be a smooth muscle cell population.Different densities of cells for seeding may be appropriate as describedherein.

In one aspect, the methods of the present invention are performed atdifferent times and for different durations following the implantationof the neo-bladder. In one embodiment, the cycling is performed on adaily basis over a period of time, on a weekly basis over a period oftime, or every other week. In another embodiment, the duration of thedaily cycling regimen is about 2 weeks, about 3 weeks, about 4 weeks,about 5 weeks, about 6 weeks, about 7 weeks, about 8 weeks, about 9weeks, about 10 weeks, about 11 weeks, about 12 weeks, about 13 weeks,about 14 weeks, or longer than 14 weeks.

In one embodiment, a daily cycling protocol for a subject may includethe steps of filling the neo-bladder for about an hour, draining thefilled neo-bladder for about an hour, and allowing the neo-bladder todrain freely, typically overnight. This protocol can be performed on dayone of the cycling regimen in the subject. This daily sequence can beperformed for a number of consecutive days after the first day. In oneembodiment, the cycling protocol may be performed on a day after day onein which the duration of the filling step is increased to about twohours, about three hours, about four hours, or more than about fourhours. In another embodiment, the filling and draining steps may berepeated more than once daily before allowing the neo-bladder to drainfreely.

In another embodiment, the subjects are catheterized post-implantation,and the cycling time is controlled by clamping and unclamping thesubject's catheter.

Those of ordinary skill in the art will appreciate that additionalcycling regimens are contemplated herein.

An example of a cycling protocol is as follows. Following implantationof a neo-bladder construct formed by seeding a neo-bladder matrix orscaffold with cells as described herein, cycling will be performed every2 weeks (14±2 day intervals) starting approximately 1 month afterimplantation and continuing until approximately Day 90. Cycling will becompleted after certain types of assessment, such as compliancemeasurement of the implanted neo-bladder, but before other types ofassessment such as fluoroscopic imaging. Cycling will be performed byre-inflating the bladder with sterile saline (warmed by incubator) afterthe completion of compliance measurement at a rate of 10-25 mL/min. Thecycling will be repeated at least 5-10 times. The starting pressure of0-10 mmHg will be achieved and recorded along with the start time. Time,volume of isotonic solution delivered, and the pressure obtained will berecorded for each cycle at the time leakage is observed around thecatheter (a.k.a. leak point), or when the volume delivered is equal tothat of the compliance measurement just performed, whichever comesfirst.

In one embodiment, the present invention provides a method of promotingregeneration of a neo-bladder implanted in a subject that includes thesteps of (a) filling the implanted neo-bladder with a fluid; (b)emptying the filled neo-bladder of step (a). In another embodiment, themethod includes step (c) repeating steps (a) and (b). In one otherembodiment, the method is commenced within the first 2 weekspost-implantation. In one embodiment, the steps (a) and (b) areperformed once daily, once weekly, or once every other week. In someother embodiments, the filling step (a) is performed for about one hourand the emptying step (b) is performed for about one hour. In yetanother embodiment, steps a) and b) are performed at least until aboutsix weeks post-implantation. In one other embodiment, steps a) and b)are not performed for more than about ten weeks post-implantation. Inanother embodiment, steps a) and b) are performed for more than aboutten weeks post-implantation. In other embodiments, the filling comprisesexpanding the neo-bladder. In another embodiment, the regenerationcomprises an increase in the capacity of the neo-bladder as compared toa neo-bladder in a subject that has not undergone cycling. In one otherembodiment, the regeneration comprises an increase in compliance of theneo-bladder as compared to a neo-bladder in a subject that has notundergone cycling. In other embodiments, the regeneration comprises anincrease in extracellular matrix development in the neo-bladder ascompared to a neo-bladder in a subject that has not undergone cycling.In one embodiment, the increase in extracellular matrix developmentcomprises the development of elastin fibers.

In one other aspect, the present invention concerns methods forproviding homeostatic regulative development of neo-bladders in mammalssuch that implanted neo-bladders are responsive to the needs of therecipient. In one embodiment, the implanted neo-bladder grows to a sizeproportionate to the recipient. In another embodiment, the methods forproviding homeostatic regulative development of a neo-bladder in asubject include the steps of (a) providing a biocompatible polymericscaffold; (b) depositing an a first cell population on or in a firstarea of said scaffold, said first cell population being substantially amuscle cell population; and (c) implanting the scaffold of step (b) intosaid subject to establish homeostatic regulative development. In oneother embodiment, the homeostatic regulative development comprisesrestoration of organ size and structure. In another embodiment, thehomeostatic regulative development comprises neo-bladder capacitiesproportionate to body weight. In one embodiment, the proportionateneo-bladder capacity is achieved at about four months post-implantation.In another embodiment, the method for providing homeostatic regulativedevelopment of a neo-bladder in a subject includes the step ofmonitoring the state of homeostatic regulative development or progressof the implanted neo-bladder. The monitoring may include a cystogramprocedure to show the position and shape of the implanted neo-bladder,and/or a measurement of urodynamic compliance and capacity.

In another aspect, the invention provides methods for prognosticevaluation of a patient following implantation of a new organ or tissuestructure. In one embodiment, the method includes the step of detectingthe level of MCP-1 expression in a test sample obtained from saidsubject; (b) determining the expression level in the test sample to thelevel of MCP-1 expression relative to a control sample (or a controlreference value); and (c) predicting regenerative prognosis of thepatient based on the determination of MCP-1 expression levels, wherein ahigher level of expression of MCP-1 in the test sample, as compared tothe control sample (or a control reference value), is prognostic forregeneration in the subject.

In another aspect, the invention provides methods for prognosticevaluation of a patient following implantation of a new organ or tissuestructure in the patient, the methods comprising: (a) obtaining apatient biological sample; and (b) detecting MCP-1 expression in thebiological sample, wherein MCP-1 expression is prognostic forregeneration in the patient. In some embodiments, increased MCP-1expression in the patient biological sample relative to a control sample(or a control reference value) is prognostic for regeneration in thesubject. In some embodiments, decreased MCP-1 expression in the patientsample relative to the control sample (or control reference value) isnot prognostic for regeneration in the subject. The patient sample maybe a test sample comprising a bodily fluid, such as blood or urine.

In some embodiments, the determining step comprises the use of asoftware program executed by a suitable processor for the purpose of (i)measuring the differential level of MCP-1 expression in a test sampleand a control; and/or (ii) analyzing the data obtained from measuringdifferential level of MCP-1 expression in a test sample and a control.Suitable software and processors are well known in the art and arecommercially available. The program may be embodied in software storedon a tangible medium such as CD-ROM, a floppy disk, a hard drive, a DVD,or a memory associated with the processor, but persons of ordinary skillin the art will readily appreciate that the entire program or partsthereof could alternatively be executed by a device other than aprocessor, and/or embodied in firmware and/or dedicated hardware in awell known manner.

Following the determining step, the measurement results, findings,diagnoses, predictions and/or treatment recommendations are typicallyrecorded and communicated to technicians, physicians and/or patients,for example. In certain embodiments, computers will be used tocommunicate such information to interested parties, such as, patientsand/or the attending physicians. In some embodiments, the assays will beperformed or the assay results analyzed in a country or jurisdictionwhich differs from the country or jurisdiction to which the results ordiagnoses are communicated.

In a preferred embodiment, a prognosis, prediction and/or treatmentrecommendation based on the level of MCP-1 expression measured in a testsubject having a differential level of MCP-1 expression is communicatedto the subject as soon as possible after the assay is completed and theprognosis and/or prediction is generated. The results and/or relatedinformation may be communicated to the subject by the subject's treatingphysician. Alternatively, the results may be communicated directly to atest subject by any means of communication, including writing,electronic forms of communication, such as email, or telephone.Communication may be facilitated by use of a computer, such as in caseof email communications. In certain embodiments, the communicationcontaining results of a prognosit test and/or conclusions drawn fromand/or treatment recommendations based on the test, may be generated anddelivered automatically to the subject using a combination of computerhardware and software which will be familiar to artisans skilled intelecommunications. One example of a healthcare-oriented communicationssystem is described in U.S. Pat. No. 6,283,761; however, the presentinvention is not limited to methods which utilize this particularcommunications system. In certain embodiments of the methods of theinvention, all or some of the method steps, including the assaying ofsamples, prognosis and/or prediction of regeneration, and communicatingof assay results or prognoses, may be carried out in diverse (e.g.,foreign) jurisdictions.

In another aspect, the prognostic methods described herein provideinformation to an interested party concerning the success of theimplantation, and the rehabilitation/treatment protocol forregeneration. In one embodiment, the methods include the steps ofdetecting the level of MCP-1 expression in a test sample obtained fromsaid subject; (b) determining the expression level in the test sample tothe level of MCP-1 expression relative to a control sample (or a controlreference value); and (c) predicting regenerative prognosis of thepatient based on the determination of MCP-1 expression levels, wherein ahigher level of expression of MCP-1 in the test sample, as compared tothe control sample (or a control reference value), is indicative of thestate of regeneration of a new organ or tissue structure.

Generally, as used herein, regeneration prognosis encompasses theforecast or prediction of any one or more of the following: developmentor improvement of a functional bladder after bladder replacement oraugmentation through implantation of a construct described herein,development of a functional urinary diversion after implantation of aconstruct described herein, development of bladder capacity or improvedbladder capacity after implantation of a construct described herein, ordevelopment of bladder compliance or improved bladder compliance afterimplantation of a construct described herein.

In all embodiments, the methods of providing a laminarily organizedluminal organ or tissue structure to a subject in need of such treatmentas described herein may include the post-implantation step of prognosticevaluation of regeneration as described above.

In all embodiments, the present invention relates to methods forproviding a new organ or tissue structure to a subject in need thatinclude certain post-implantation monitoring steps. In one embodiment,the effect and performance of an implanted constructs is monitored, suchas through ultrasound imaging, pyelogram, as well as urine and bloodanalysis at different time-points after implantation. These monitoringsteps are described in further detail in Examples 3-6.

7. Kits

The instant invention further includes kits comprising the polymericmatrices and scaffolds of the invention and related materials, and/orcell culture media and instructions for use. The instructions for usemay contain, for example, instructions for culture of the cells oradministration of the cells and/or cell products. The instructions foruse may also contain instructions for pre-treating, folding or otherwisepreparing the polymeric matrices and scaffolds of the invention forlaparoscopic implantation.

In one embodiment, the present invention provides a kit comprising ascaffold as described herein and instructions. In another embodiment,the scaffold of the kit is one or more of the following: a bladderaugmentation scaffold, a bladder replacement scaffold, a urinary conduitscaffold, or a muscle equivalent scaffold.

8. Reports

The methods of this invention, when practiced for commercial purposesgenerally produce a report or summary of the regenerative prognosis. Themethods of this invention will produce a report comprising a predictionof the probable course or outcome of regeneration before and after anysurgical procedure to provide a construct described herein. The reportmay comprise information on any indicator pertinent to the prognosis.The methods and reports of this invention can further include storingthe report in a database. Alternatively, the method can further create arecord in a database for the subject and populate the record with data.In one embodiment the report is a paper report, in another embodimentthe report is an auditory report, in another embodiment the report is anelectronic record. It is contemplated that the report is provided to aphysician and/or the patient. The receiving of the report can furtherinclude establishing a network connection to a server computer thatincludes the data and report and requesting the data and report from theserver computer. The methods provided by the present invention may alsobe automated in whole or in part.

The following examples are offered for illustrative purposes only, andare not intended to limit the scope of the present invention in any way.

All patent, patent applications, and literature references cited in thepresent specification are hereby incorporated by reference in theirentirety.

EXAMPLES Example 1 Peripheral Blood and Adipose Tissue as a Source ofSMCs

Blood-Derived Cells

Smooth muscle cells have been successfully isolated from canine,porcine, and human peripheral blood. Briefly, a dilution of 50 ml ofperipheral blood 1:1 with phosphate buffered saline (PBS; 100 mL finalvolume) was prepared and layered onto Histopaque, a density gradientmaterial, and centrifuged at 1,354×g for 20 minutes at room temperature.After centrifugation, four layers will be clearly defined in the densitygradient (from top to bottom): serum, buffy coat, Histopaque, red bloodcells. The mononuclear cells are located in the buffy coat, whichappears as an opaque white/gray band. The buffy coat was withdrawn andtransferred into a separate 50 ml conical tube. Dilute to 50 mL withPBS. Centrifuge the samples at 711×g for 10 minutes at room temperatureto pellet cells. Resuspend pellet and culture the cells. Whenappropriate cell numbers are reached by subsequent cell passaging, analiquote is fixed and processed for end-point analysis includingimmunodetection of expressed smooth muscle cell proteins, nucleic aciddetection of smooth muscle cell mRNA transcripts, cellular contraction,cytokine and enzyme synthesis.

Results

Media selection. The mononuclear fraction of a single 40-50 mL canineperipheral blood sample was resuspended in six different mediaformulations and seeded into 6-well Primaria or collagen-coated plates.

As shown in FIG. 14A-E, after one week of culture, small adherentcolonies and small cell aggregates were observed in all conditions (DMEMmedia isolations are not shown) but the identity of the cell types wereindiscernible. Small clusters and cell aggregates were observed onPrimaria culture dishes when grown in alpha-MEM+10% FBS, EGM-2 mediumwith all of the accompanying supplements, and EGM-2 with selectedaccompanying supplements (minus VEGF and FGF2) (A, C, E) and collagentype I coated on tissue culture plastic plates grown in the same medias(B, D, F). Similar results were seen in peripheral blood cultures grownin DMEM formulations (data not shown).

As shown in FIG. 15, after two weeks of culture, outgrowth colonies andsmall monolayers were observed in alpha-MEM on Primaria (left panels)and collagen-coated plates (middle panels). Morphologically, thesecolonies appeared smooth muscle (FIG. 15, top panels) or endothelial(FIG. 15, middle panels). Outgrowth colonies of smooth muscle (FIG. 15,top panels) or endothelial (FIG. 15, middle panels) morphology alsoformed in other media/substrate conditions (right panels). Somemacrophages were initially maintained in alpha-MEM (FIG. 15, bottom leftand middle panels), but did not carry over into subsequent passages.Cells isolated in αMEM with 10% FBS on Primaria plates were of smoothmuscle (top left panel) or macrophage (bottom left panel) morphology. Noendothelial cells were isolated under these conditions (middle leftpanel). Cells isolated in αMEM/10% FBS on collagen I plates were ofsmooth muscle (top middle panel), endothelial (middle middle panel) andmacrophage (bottom middle panel) morphology. Other media/substrateformulations such as EGM-2 (top right panel) and DMEM supplemented with20% FBS (middle right panel) also permitted outgrowth of mescenchymal-and endothelial-like cells.

Of the twelve media/substrate conditions, Primaria plates withalpha-MEM/10% FBS contained the most homogeneous isolation of smoothmuscle cells (FIG. 15, top left panel) without outgrowth colonies ofendothelial cells. Cells isolated on Primaria plates and expanded onNunclon surfaces (in alpha-MEM/10% FBS) exhibited the classical ‘hilland valley’ morphology that is typical of smooth muscle cells (SMC), andis consistent with descriptions in other studies (Kassis et al. (2006);Koerner et al. (2006); Simper et al. (2002), supra).

As shown in FIG. 16, these cells also maintained this morphology forseveral passages (FIG. 16A-G). Images of porcine carotid artery SMC(FIG. 16H) and dog bladder SMC (FIG. 16I) are shown for comparison. Thesmooth muscle cells at later passages (FIG. 16F, G) became larger andmore spread out. Early passages (A-E) resemble smooth muscle cells (SMC)isolated from porcine carotid artery (H) and dog bladder (I). Laterpassages of smooth muscle cells (F, G) are larger and more spread out,suggesting a smooth muscle phenotype.

Adipose-Derived Cells

Smooth muscle cells have been isolated from porcine adipose tissueaccording the following procedure. All procedures are performed in thebiosafety hoods.

Obtain adipose sample. Store at room temperature or 4° C. for no morethan 24 hours prior to use in biosafety container.

Prepare collagenase solution by adding 1 gm of BSA and 0.1 to 0.3 gm ofcollagenase per 100 ml of PBS. Filter the solution through a 0.2 μmfilter unit. Warm to 37° C.

Add equivalent volume of Collagenase solution per adipose volume to eachcentrifuge bottle. One tissue volume of collagenase solution is required(i.e. 10 ml of collagenase solution per 10 ml adipose tissue).

Wipe the tubes with disinfectant, cap, wrap with parafilm and place in a37° C. incubator on a rocker for 60 minutes. Alternatively, tubes may beplaced in a 37° C. water bath and vigorously shaken by hand every 20min.

Centrifuge at 300×g at Room Temperature for 5 minutes.

Take the tubes out of the centrifuge and shake them vigorously for 10seconds to thoroughly mix the cells. This is to complete the separationof stromal cells from the primary adipocytes.

Centrifuge again at 300×g for 5 minutes. Carefully aspirate off the oilon top, the primary adipocytes (yellow layer of floating cells), and thecollagenase solution. Leave behind approximately 10 ml of the browncollagenase solution above the pellet so that the stromal-vascularfraction (dark red cells on bottom) is not disturbed.

Resuspend the pellet of cells in PBS with 1% BSA and filter usingSteri-Flip.

Centrifuge the cells at 300×g for 5 minutes and aspirate the remainingcollagenase solution. When aspirating, the tip of the pipette shouldaspirate from the top so that the oil is removed as thoroughly aspossible. The cell pellet should be tightly packed at the bottom.

Add 10 ml of tissue culture medium to each centrifuge tube and resuspendthe cells. Pool the cells to one tube and centrifuge again.

Aspirate supernatant. Suspend the cells in 10 ml of medium.

Divide the cells equally and accordingly to the appropriate number offlasks. 24-72 hours after plating, aspirate medium from flask. Wash withPBS and aspirate.

Add the original volume per flask of fresh medium.

Cells will be grown to 80-90% confluence and then either passaged orcryopreserved.

When appropriate cell numbers are reached by subsequent cell passaging,an aliquot is fixed and processed for immunodetection of expressedsmooth muscle cell proteins.

FIG. 17 concerns the morphological assessment of cultures. Themorphology was assessed after 3 to 5 days in culture. Human and porcinecells derived from adipose tissue exhibit smooth muscle cellmorphological characteristics (FIG. 17). The cells demonstrate ahill-and-valley morphology and exhibit additional characteristics suchas spindly shaped, flattened and fibroblast-like upon passage, elongatedand arranged in parallel rows, and a “whirled” appearance of growth, allof which are typical of cultured smooth muscle cells.

Smooth muscle markers. Increased expression of contractile genes (andthe proteins they encode) is associated with SMC maturation (Jeon et al.J Cell Sci 119, 4994-5005 (2006); Ross et al. J Clin Inves. 116,3139-3149 (2006); Sinha et al. Am J Physiol Cell Physiol 287, 1560-1568(2004)). Myocardin is a transcriptional regulator of genes that encodesmooth muscle contractile proteins, among which include SM22, alphasmooth muscle actin, smooth muscle myosin heavy chain, and calponin (Qiuet al. (2005) Circ Res 983-991; Wang et al. (2003) Proc Natl Acad Sci100:7129-7134; Yoshida et al. (2003) Circ Res 92:856-864). Myocardin isrequired for smooth muscle differentiation, and is sufficient to drivesmooth muscle gene expression in some cell types (Milyaysky et al.(2007) Cancer Cell 11:133-146; van Tuyn et al. (2005) supra; Wang et al.(2003), supra; Yoshida et al. (2003), supra). We determined if thesmooth muscle cells isolated from blood or adipose tissue expressed thesmooth muscle cell markers myocardin, smooth muscle alpha actin, SM22,myosin heavy chain, and calponin by isolating total RNA and performingsemi-quantitative RT-PCR.

As shown in FIG. 18, the results indicate that these cells express allof these smooth muscle cell markers at the gene level, consistent withthe smooth muscle cell markers found in bladder smooth muscle cells.These data support the notion that these smooth muscle cells isolatedfrom peripheral blood or adipose tissue have properties of smooth musclecells.

Phenotypic characterization. We have already shown that these peripheralblood isolated smooth muscle cells express a transcriptional regulatorof smooth muscle gene expression as well as specific smooth musclecontractile proteins (FIG. 18). FIG. 18 shows RT-PCR analysis for geneexpression of SMC markers myocardin, smooth muscle alpha-actin, SM22,smooth muscle myosin heavy chain, and calponin Samples were from smoothmuscle cells isolated from porcine adipose, peripheral blood, andbladder (passage 4). The SMCs isolated from adipose tissue can becultured 3-5 days between each passage, while the SMCs isolated fromblood can be cultured for 14 days before the first passage and then 3-5days for additional passage. Gene expression for beta-actin was used asan internal loading control for the gel. Expression profiles for adiposeand peripheral blood cell isolates are comparable to that of the bladderSMC.

FIG. 19 shows immunofluorescence staining that was performed utilizing avariety of antibodies directed towards smooth muscle cell expressedprotein markers. The markers alpha-actin, SM22, calponin, and smoothmuscle myosin heavy chain were examined in smooth muscle cells isolatedfrom porcine adipose, peripheral blood, and bladder. These proteins areall involved in the contractile function of smooth muscle cells. Smoothmuscle cells at multiple passages stained positively for smooth musclealpha actin, SM22, calponin, and smooth muscle myosin heavy chain.Subcellular localization of these proteins was virtually identical insmooth muscle cells compared to bladder SMC. Detailed staining of theseproteins in the stress fibers of the cells was noted. This pattern ofstaining is typical and expected for smooth muscle cells.

FIG. 20 shows immunostaining of smooth muscle cells isolated from humanperipheral blood (passage 5). Probes for smooth muscle alpha actin,SM22, and calponin were used. Dual staining for smooth muscle alphaactin and calponin (top right panel) reveals co-expression of these twoproteins within the same cells. This simultaneous expression of morethan one smooth muscle cell marker in a single cell further supports thenotion that these smooth muscle cells.

Contractility. Since the peripheral blood derived smooth muscle cellsexpress smooth muscle contractile proteins, we performed athree-dimensional gel contraction assay to assess their contractilefunction. SMC have been shown to spontaneously induce contraction of acollagen matrix when embedded in a three-dimensional gel (Travis et al.(2001) Circ Res 88:77-83). Adipose tissue-derived smooth muscle cellswere also tested for contractility.

FIG. 21 shows that porcine blood-derived (A) and porcine adiposetissue-derived (B) cells contract to a degree comparable to that ofbladder smooth muscle cells (C). The addition of EDTA to the mixtureinhibits contraction, thus supporting the idea that the contraction iscalcium dependent, another characteristic of smooth muscle cells. Thesedata indicate that diameter reduction is dependent on contractile cells,and that the cells function in this capacity. The cells were seeded at500,000 cells/ml and found to be capable of contraction as demonstratedby a reduction of collagen gel diameter after two days. Porcine bladdersmooth muscle cells were used as a positive control. To demonstrate thecalcium dependence of this contraction, the calcium chelator EDTA wasadded to separate samples to inhibit contraction. These results confirmthe ability of the cells to contract in a calcium-dependent mannersimilarly to bladder-derived smooth muscle cells.

Growth kinetics. In order to utilize smooth muscle cells in cell therapyapplications, it is important to determine if the required cell numberscan be achieved in an acceptable time frame. The results from canine andporcine studies indicate that smooth muscle colonies (from a 40 mlsample of peripheral blood) can be observed as early as 7 days postseeding, and can readily be passed within 14 days (FIGS. 14 and 15). Inone study, 1.2 million cells were obtained after 18 days of culture (endof passage 2), at which time they were cryopreserved. These particularcells were thawed ˜50 days later, and routinely passed when ˜80%confluent to determine growth kinetics. Six days after thawing, the cellpopulation expanded to 16.7 million cells (end of passage 3). Afteranother 7 days of culture, the cell population reached 31.7 millioncells (end of passage 4). This initial study indicates that 30 millioncells can be achieved in roughly 30 days of culture.

FIG. 22 concerns the limited proliferation potential of the cells. FIG.22 shows the growth of smooth muscle cells isolated from human adiposetissue as a function of the numbers of cells recovered per unit area.These data indicate that between passages 4 and 5, the number ofrecovered cells begins to decline, supporting the contention that thesecells have a limited and finite proliferative capacity, which ischaracteristic of progenitor cells, but not true stem cells.

FIG. 23 shows the growth of smooth muscle cells isolated from porcineadipose, peripheral blood, and bladder smooth muscle as a function ofthe number of recovered cells per passage. As illustrated, dramaticexpansion in cell numbers is achieved between passages 2 and 3, over atime frame of 2-4 weeks, enabling recovery of tens of millions of cells.This demonstrates the limited or finite proliferation potential of theadipose-derived cells.

Contact inhibition of proliferation. The smooth muscle cells isolatedfrom peripheral blood and adipose tissue exhibit contact inhibition ofproliferation. For example, the morphological assessment of these cellsprovided in FIGS. 14-17 demonstrates the presence of contact inhibitionof proliferation over several passages. The cells do not continueproliferating upon contact with each other. In contrast, MSCs do notexhibit contact inhibition of proliferation and they can be observedpiling on top of each other, similar to foci formation in transformedcell cultures. For example, Zhou et al. report on the isolation andculturing of MSCs from the mononuclear cell fraction of mouse bonemarrow, and observe that after three passages the cultured MSCsdemonstrated a loss of contact inhibition (see page 10850 and FIG. 1A)(Cancer Res. 2006; 66(22):10849-10854).

Cytokine MCP-1 production. MCP-1 is a normal product of bladder detrusorcells. In aortic smooth muscle cells, MCP-1 plays a role inregeneration. MCP-1 is best known for its ability to recruit mononuclearcells. It is however more than a chemokine; it is also a potent mitogenfor vascular smooth muscle cell proliferation and recruits circulatingmonocytes to the area of vessel injury. Monocytes are typicallytransformed to macrophages which can serve as reservoirs for cytokinesand growth factors. Macrophages and muscle precursor cells are bothtargets for MCP-1 signaling. This cytokine has been implicated in stemand progenitor cell recruiting within the body, potentially contributingto the regenerative process.

In order to quantitate MCP-1 produced by human peripheral blood smoothmuscle cells, an ELISA based assay system from R&D Systems was employed.Medium samples were assayed in duplicate and compared to a standardcurve to provide estimated MCP-1 levels and reported as ug/24 hr/onemillion cells. Expression of the cytokine MCP-1 for cells isolated fromhuman bladder smooth muscle, adipose, peripheral blood, and bladderurothelium (negative control) was determined.

FIG. 24 shows the results from this analysis indicates that humanperipheral blood-derived and human adipose tissue-derived smooth musclecells produce MCP-1 at levels comparable to that of human bladder smoothmuscle cells. These data support the conclusion that, just like bladderSMC, MCP-1 is expressed by the smooth muscle cells isolated from adiposeand peripheral blood. In addition, these data lead us to hypothesizethat the production of MCP-1 may play a critical role in regeneration bydirectly or indirectly causing muscle progenitor cells to berecruited/migrate or to proliferate within the construct.

Discussion. Isolated smooth muscle cells from adipose demonstrateseveral smooth muscle cell characteristics. Our studies have indicatedthat the cells can readily be isolated from adipose using standardenzymatic digestion and low-speed centrifugation protocols. Cells can beexpanded very rapidly, perhaps reaching ˜30 million cells within amonth's time. Our studies have further demonstrated that these cellsmay, in fact, represent a smooth muscle cell population rather than atrue stem cell population, as smooth muscle markers are present as earlyas passage 3. Expression of SMC marker mRNA can be observed as early asP0, as demonstrated by RTPCR. Furthermore, the smooth muscle cellsisolated from are capable of contractile function as demonstrated bystandard collagen gel contraction assays.

Characterization of smooth muscle cells. We have already shown thatduring subsequent passages, the smooth muscle cell cellular morphologyis retained. There is also good correlation of smooth muscle markers atboth the gene and protein levels.

Cytokine induction. Expression of MCP-1 by adipose smooth muscle cellshas lead us to hypothesize that the production of MCP-1 may play acritical role in neo-organ or tissue structure regeneration by directlyor indirectly causing native mesenchymal stem cells to berecruited/migrate or to proliferate within the construct.

Example 2 MCP-1 Production and Cell Density

Conditioned medium from cultures of bladder smooth muscle cells wereanalyzed using commercially available kits for the detection andquantitation of MCP-1. Conditioned media samples from 9 constructs (3from each of 3 seeding levels) and the paired SMC cells used for seedingthe constructs were tested for MCP-1 levels. The results are shown inTable 2.1.

TABLE 2.1 Sam- Test ple cIL2 cIL6 cIL10 cMCP-1 cIFNg cTNFa cTGFb ID IDpg/ml pg/ml pg/ml pg/ml pg/ml pg/ml pg/ml 1 TT1 <1.0 <9.8 1.0 <3.7 <2.4<0.2 2 TT2 <1.0 8.8 <2.0 39.6 <2.4 <0.2

In order to quantitate MCP-1 present in the construct medium, an ELISAbased assay system specific for Canine MCP-1 from R&D Systems wasemployed. Samples were assayed in duplicate and compared to a standardcurve to provide estimated MCP-1 levels in construct medium. As shown inFIG. 25, the results from this analysis show a positive correlationbetween MCP-1 production and the density of cells seeded. Table 2.2shows MCP-1 quantitation of construct medium as determined by R&DSystems ELISA.

TABLE 2.2 MCP-1 Group Std Group Construct pg/ml Average Dev  4 million1151 71 65 24 1152 102 1153 59 1154 80 1155 74 1156 24 1157 70 1158 3912 million 1159 253 188 135 1160 85 1161 412 1162 167 1163 69 1164 3491165 91 1166 78 25 million 1167 183 385 207 1168 307 1169 181 1170 5271171 771 1172 534 1173 260 1174 321

Table 2.3 shows a comparison of the average MCP-1 levels from each groupin which it can been seen that the resulting ratios parallel thedifferences in seeding densities.

TABLE 2.3 4 million 12 million Average Cell Cell 25 million Group MCP-1MCP-1 # MCP-1 # MCP-1 Cell #  4 million 65 0.35 0.33 0.17 0.16 12million 188 2.89 3.00 0.49 0.48 25 million 385 5.92 6.25 2.05 2.08

Results indicated that there was a positive correlation between cellnumber and MCP-1 levels detected in the media. It had been previouslynoted that some tissue from a regenerated canine bladder (approximately9 million cells seeded) processed for SMC explantation contained morefat than is typically observed in native and regenerated canine tissue.The tissue when explanted was very soft and the explants when viewedcontained fatty tissue in greater proportion to that observed withnative tissue. The media on these explant plates also exhibited a“sheen” on the surface typically observed when fatty tissue is present.These observations suggest a role for MCP-1/CCR-2 interaction in fatdeposition/adipogenesis of regenerated bladder tissue.

Example 3 Implantation of Neo-Urinary Conduit Constructs in Swine

The objective of this study was to investigate the surgical implantationof the neo-urinary conduit and evaluation of the post surgical care, aswell as to assess the regeneration of urinary-like tissue followingimplantation of the Neo-Urinary Conduit (NUC) test articles and theability of swine peritoneum to provide a vascular supply andwater-tightness to the implant.

Neo-Urinary Conduit (NUC) construct test articles were comprised of ascaffold formed from nonwoven polygycolic acid (PGA) felts andpoly(lactic-co-glycolic acid) polymers (PLGA) with or without autologoussmooth muscle cells (SMC). Cells previously removed from an animal wereused to produce the construct that was implanted in the same animal. Aconstruct refers to the sterile tube-shaped biomaterial comprised of ascaffold formed from nonwoven polygycolic acid (PGA) felts andpoly(lactic-co-glycolic acid) polymers (PLGA) combined with autologousSMC. The term scaffold-only refers to the sterile tube-shapedbiomaterial comprised of a scaffold formed from nonwoven polygycolicacid (PGA) felts and poly(lactic-co-glycolic acid) polymers (PLGA)without cells.

In this study, the constructs used correspond to test articles comprisedof scaffold and SMC and scaffold only refers to a test article comprisedof scaffold without SMC. Seven female Gottingen minipigs were dividedinto three groups: N=1 in Group 1 (scaffold-only), N=3 in Group 2(scaffold seeded with blood derived SMC) and N=3 in Group 3 (scaffoldseeded with adipose derived SMC) for implantation with the testarticles.

Swine were considered a suitable animal model for evaluation of theNeo-Urinary Conduit given the similarities between swine and humanabdominal and upper urinary tract anatomy, surgical manipulationstrategies, stoma placement and healing, postsurgical care, andperitoneal anatomy. Swine is also a well established animal model ofwound healing in skin, closely approximating the normal process ofhealing in humans, allowing evaluation of stoma healing. Gottingenminipigs were chosen as the breed based on their slow average growthrate during the 3-month study duration. Autologous SMCs were obtainedfrom adipose tissue biopsies and venous blood samples from all animalsapproximately 10-11 weeks prior to test article implantation. Specifiedtest articles were surgically implanted on Day 0 in each group. Aftersurgical removal of the bladder (total cystectomy) the ureters werestented and mobilized for anastomosis to the inflow (cranial) end of thetest article. Parietal peritoneum was separated from the abdominal wallstarting from the linea alba at midline and bilaterally towards theright and left side of the abdominal wall. The peritoneum was transectedon the left side and used to wrap the implants towards the right ofmidline portion which provided the vascular source and a watertighturine channel, and formed a tubular connection (atrium) between thecaudal end of the implant (located in the intra-abdominal cavity) andthe skin. The implant's caudal end terminated within the peritonealatrium approximately 5 to 7 cm away from the skin stoma. The atrium wasextended using the cranial peritoneal wrap which traversed the abdominalwall and exited the skin near the xiphoid (off midline, right side). Theexternalized peritoneum was sutured to the skin to form aperitoneum-cutaneous junction and peritoneal-lined stoma lumen.

The suture strands that were connected to the ureteral stents wereexteriorized through the stoma for future removal. The abdominalincision was closed with non-absorbable Prolene suture. The skin wasclosed in a routine fashion. A Foley catheter was inserted into thestoma to allow urine passage during stoma healing. The same surgicalprocedure was used for all animals Following removal of the Foleycatheter, all animals were fitted with TRACOE® stoma ports to facilitateurine drainage. The animals were able to dislodge the stoma port, so an8Fr Foley catheter was used to aid urine drainage. Detritus buildup inthe atrium and stoma led to the use of a larger diameter modifiedextension set (study specific) to manage the stoma.

Stoma maintenance and replacement was scheduled weekly and was done onas needed basis. Blood samples were collected, analyzed and the resultsrecorded at baseline, weekly during weeks 1 through 4 post-implantation,week 8, and necropsy for hematology and serum chemistry. Urine sampleswere collected, analyzed and the results recorded at baseline andnecropsy for urinalysis. Imaging (fluoroscopy, ultrasonography, and/orendoscopy) of the constructs, ureters, and kidneys was performed atweeks 2, 4, 8 and necropsy during the study. Imaging was also performedas needed in response to adverse clinical signs (e.g. observed lack ofurine flow or suspected fistula formation). A fistula refers to anabnormal duct or passage that connects an abscess, cavity or holloworgan to the body surface or to another hollow organ (for example,between intestines or between the intestines and conduit).

At necropsy, the abdominal cavity was opened, the conduit visualized andphotographed before the conduit was removed en bloc with stoma, kidneysand ureters. Representative tissue samples of the entire urinary tractfrom kidneys to skin stoma, regional lymph nodes, and any other lesionsobserved grossly were obtained. All tissue samples were placed in 10%Neutral Buffered Formalin (NBF) for histological processing andevaluation. Post fixation, tissues were processed routinely tomicroslides and stained with hematoxylin and eosin (H&E) and Masson'strichrome. Slides were evaluated microscopically.

Results: Implantation (Surgical Methodology): All animals were recoveredfrom implantation surgery uneventfully and the stoma was visualized tobe draining urine. The animal model was considered appropriate forevaluating the surgical procedures for implanting the NUC (Neo-UrinaryConduit).

Morbidity and Mortality: Animals survived 28-83 days. One of 7 animalssurvived until scheduled sacrifice (Animal 4 of Group 2, 83 days). Sixof 7 animals were sacrificed unscheduled: animal 5 of Group 3 waselectively euthanized 28 days post-implantation for histopathologicalanalysis and 5 animals were euthanized for poor clinical conditionbetween 38 and 63 days post-implantation. (animal 1 of Group 1, animals2 and 3 of Group 2, and animals 6 and 7 of Group 3). These unscheduleddeaths occurred in all treatment groups and were attributed to viralinfection and/or obstruction-related pathology with damage to the upperurinary tract.

Evidence of Porcine Circovirus Type-2 (PCV-2 evidence gathered duringhistopathology of harvested tissues) infection was observed in 3/7animals. Two animals with PCV-2 infection were unscheduled deathanimals. These included animal 2 of Group 2 euthanized on day 38 andanimal 7 of Group 3 euthanized on day 63. The third animal identifiedwith PCV-2 infection (animal 4 of Group 2), survived to scheduledsacrifice (83 days). Obstruction of urine flow through the conduit andstoma contributed to the morbidity in 4/6 unscheduled deaths animals.These included animal 1 of Group 1 euthanized on day 47; animals 2 and 3of Group 2, euthanized on days 38 and 40; and animal 6 of Group 3euthanized on day 39.

Post-Surgical Care: Day 1-30 post-implantation: All animals requiredpostoperative stoma management (e.g. flushing and cleaning of debris andreplacement of catheter or stoma port when dislodged weekly and on asneeded basis) regardless of treatment. Debris is formed during thehealing and regenerative processes. Exfoliated tissue cells,inflammatory exudate and scaffold biodegradation are sources of debris.Without proper outflow (e.g., with obstruction), the stagnated debrisforms a detritus: a semisolid bolus within the lumen of the conduit.

Loss of appetite and lethargy were observed in all animals. Thesepost-operative procedures and clinical signs were not considereduncommon following abdominal surgery in swine. Based on the first30-days post-implantation, none of the post-operative manipulations orfindings were considered sufficiently considerable at the time ofevaluation to change the established surgical procedures orpost-surgical care.

Day 31 post-implantation to necropsy: All animals demonstrated partialor complete obstruction of the urine outflow. Obstruction of urine flowwas observed with or without debris accumulation. Ventral abdomenpositioning of surgically implanted test article contributed to physicalobstruction of urine flow in the quadruped animal model where the weightof the overlying abdominal organs contributed to conduit closure,adhesion and fistula formation, and secondary upper urinary tract renalcomplications (e.g., dilation, inflammation, and/or infection of uretersor kidney). Adhesion refers to the union of two tissue surfaces.Intra-abdominal and/or pelvic adhesions are common post-surgicalcomplications. At necropsy, adhesions to conduit or ureters wereobserved grossly and radiographically, and efforts for microscopiccorrelates were attempted.

In addition, the urine flow obstruction was exacerbated by the use ofperitoneum to form the atrium, causing partial or full urinaryobstruction with subsequent detritus build-up and bacterial infection.The atrium refers to the anterior connecting chamber that allowed forurine passage through the abdominal wall. This segment was made by themost anterior tube-like portion of the peritoneal wrap connecting thecaudal end of the test article (located in the intra-abdominal cavityapproximately 5 to 7 cm away from the skin) to the skin.

Surgical placement of the test article was the same in all studyanimals; therefore, obstruction-related complications had a similarpathobiological mechanism in all groups (i.e. abdominal content pressureand a peritoneal atrium contributing to detritus build up).

Regeneration: Regeneration of urinary-like tissue was evident as earlyas day 28, with presence of urothelium, lamina propria and smooth musclebundles at the ureter-conduit junction (UCJ) in an electively euthanizedanimal 5 of Group 3(adipose-derived SMC). The regenerative process atthe ureteral end of the implant resulted in urinary-like tissueformation that was comparable among animals receiving a constructimplant (Groups 2 and 3). The extent of urinary-like tissue regenerationin the construct groups (Groups 2 and 3) was influenced by duration ofanimal survival post-implantation. The one animal surviving to scheduledsacrifice (animal 4 of Group 2 at day 83) had urothelium and smoothmuscle present in the UCJ, cranial and mid portions of the conduit inspite of a detected viral infection. However, the peritoneum-only atriumwas insufficient to support urinary-like tissue regeneration and thetissue formed in the atrium had a wall comprised of fibrous connectivetissue without urothelial mucosal lining The point of transition fromconduit to atrium varied between animals because the caudal end of theimplant floated freely within the peritoneal wrapping making thetransition from conduit to atrium difficult to define at necropsy. Thetypical composition of the (presumed) caudal conduit was organizedcollagen with associated fibroblasts and/or myofibroblasts. Peritoneumatrium appears to be insufficient for urinary-like tissue regeneration,but the peritoneum does serve as a source of vascularization to NUCimplants.

Conclusions: The swine animal model proved appropriate for evaluatingthe surgical application of the Neo-Urinary Conduit in this pilot studybecause all animals recovered from surgery and urinary diversion wasachieved. In addition, the swine model was appropriate for evaluatingpost-operative care of urinary flow obstruction and its impact to theupper urinary tract. Finally, the swine model was appropriate forevaluating the ability of the test articles to regenerate urinary-liketissue in an environment complicated by detritus accumulation andbacterial colonization, viral infection, and enteric adhesions andfistulas. The surgical methodology was determined to be successfulalthough anatomical placement of the urinary diversion on the ventralabdominal floor of a quadrupedal animal resulted in partial obstructionof urine outflow. The animal model was considered appropriate forevaluating the surgical application, postsurgical care and functionalityof the Neo-Urinary Conduit.

Post-surgical findings during the first 30 days following implantsurgery revealed findings that were not considered uncommon followingurinary diversion surgery in the pig.

Although several confounding factors occurred during the study (i.e.surgical placement on the ventral abdominal floor, use of peritonealatrium and viral infection), regeneration of urinary-like tissue wasevident as early as day 28, with presence of urothelium, lamina propriaand smooth muscle bundles at the ureterconduit junction in an electivelyeuthanized animal (animal 5 of Group 3, adipose derived SMC).

The extent of urinary-like tissue regeneration in the construct groups(Groups 2 and 3) was influenced by duration of animal survivalpost-implantation. The one animal surviving to scheduled sacrifice(animal 4 of Group 2 at Day 83) had urothelium and smooth muscle presentin the UCJ, cranial and mid portions of the conduit in spite of adetected viral infection.

There were no apparent differences observed in the regenerative processwhen scaffolds were seeded with SMC derived from blood or adipose(Groups 2 and 3, respectively) suggesting equivalence between SMCsources in promoting regeneration.

The tissue formed from peritoneum in the atrium segment of the conduithad a wall comprised of fibrous connective tissue without urotheliallining.

Experimental Design

Overview: Seven female Gottingen minipigs were divided into threegroups: N=1 in Group 1 (scaffold only), N=3 in Group 2 (blood-derivedSMC), and N=3 in Group 3 (adipose-derived SMC) and implanted with thetest articles. Autologous SMC were obtained from adipose tissue biopsiesand venous blood samples from all animals approximately 10-11 weeksprior to test article implantation. Specified test articles weresurgically implanted on Day 0 in each group. After surgical removal ofthe bladder (total cystectomy) the ureters were stented and mobilizedfor anastomosis to the inflow (cranial) end of the test article.Parietal peritoneum was separated from the abdominal wall starting fromthe linea alba at midline and bilaterally towards the right and leftside of the abdominal wall. The peritoneum was transected on the leftside and used to wrap the implants towards the right of midline whichprovided the vascular source and a watertight urine channel, and formeda tubular connection (atrium) between the caudal end of the implant(located in the intra-abdominal cavity) and the skin. The implant'scaudal end terminated within the peritoneal atrium approximately 5 to 7cm away from the skin stoma The atrium was extended using the cranialperitoneal wrap which traversed the abdominal wall and exited the skinnear the xiphoid (off midline, right side). The externalized peritoneumwas sutured to the skin to form a peritoneum-cutaneous junction andperitoneal-lined stoma lumen. The suture strands that were connected tothe ureteral stents were exteriorized through the stoma for futureremoval. The abdominal incision was closed with non-absorbable Prolenesuture. The skin was closed in a routine fashion. A Foley catheter wasinserted into the stoma to allow urine passage during stoma healing. Thesame surgical procedure was used for all animals.

Following removal of the Foley catheter, all animals were fitted withTRACOE® stoma ports to facilitate urine drainage. The animals were ableto dislodge the stoma port, so an 8Fr Foley catheter was used to aidurine drainage. Detritus buildup in the atrium and stoma led to the useof a larger diameter modified extension set (study specific) to managethe stoma. Stoma maintenance and port/catheter replacement was scheduledweekly and was done on as needed basis.

Blood samples were collected, analyzed and the results recorded atbaseline, weekly during weeks 1 through 4 post-implantation, week 8, andnecropsy for hematology and serum chemistry. Urine samples werecollected, analyzed and the results recorded at baseline and necropsyfor urinalysis. Imaging (fluoroscopy, ultrasonography, and/or endoscopy)of the constructs, ureters, and kidneys was performed at weeks 2, 4, 8and necropsy during the study. Imaging was also performed as needed inresponse to adverse clinical signs (e.g. observed lack of urine flow orsuspected fistula formation). At necropsy, the abdominal cavity wasopened, the conduit visualized and photographed before the conduit wasremoved en bloc with stoma, kidneys and ureters. Representative tissuesamples of the entire urinary tract from kidneys to skin stoma, regionallymph nodes, and any other lesions observed grossly were obtained. Alltissue samples were placed in 10% Neutral Buffered Formalin (NBF) for24-48 hours prior to shipping to Vet Path Services, Inc. forhistological processing and evaluation. Post fixation, tissues wereprocessed routinely to microslides and stained with hematoxylin andeosin (H&E) and Masson's trichrome. Slides were evaluatedmicroscopically. The pathology report appears in the Example below.

The following table 3.1 provides a summary of the study design.

TABLE 3.1 Biopsy Estimated Group No. of Procedure Surgical PostoperativeNecropsy No. Treatment Animals (~Day 70) Procedure (Day 0) proceduresTime Point 1 Scaffold only 1 Removal of Cystectomy Fluoroscopic and 84 ±5 days 2 Autologous 3 adipose followed by neo- ultrasonic Blood SMCbiopsy, blood urinary conduit examination, collection implantation withgeneral health transposition of assessment and ureters to inflowclinical treatment end. Whole test as necessary, article wrapped inclinical pathology peritoneum, with a and urinalysis peritonealtransition from end of test article through skin to create incontinentstoma. 3 Autologous 3 adipose SMC

Swine were considered as the optimal animal model for evaluation of theNeo-Urinary Conduit given the similarities between swine and humanabdominal and upper urinary tract anatomy, surgical manipulationstrategies, stoma placement and healing, and post-surgical care. Swineis a well-established animal model of wound healing in skin, closelyapproximating the normal process of healing in humans, allowingevaluation of stoma healing. The omentum was previously validated forproviding a blood supply and a water tight surface for urinary tissueregeneration in dogs. In the current studies, the ability of peritoneumto provide a vascular supply and watertightness for the NUC was beingevaluated and swine is the only large animal species with a parietalperitoneum similar to that of humans. Gottingen minipigs were chosen asthe breed of swine based on a slow average growth rate during the3-month study duration.

Materials and Methods

Test Devices—The test articles were i) a conduit shaped scaffoldcomprising a synthetic lactide/glycolide polymer seeded with autologousadipose-derived pig smooth muscle cells (2.5×10⁷ cells or 2.5×10⁷cells); ii) a conduit shaped scaffold comprising a syntheticlactide/glycolide polymer seeded with autologous blood-derived pigsmooth muscle cells (2.5×10⁷ cells or 2.5×10⁷ cells) and iii) a conduitshaped scaffold comprising a synthetic lactide/glycolide polymer withoutany cells seeded.

Animals. A total of 7 animals were implanted with the test articles. Theanimals were initially housed in individual cages. After test articleimplantation, the animals were transferred into a steel pen with twopigs in each pen. The animals were fed twice daily in the morning and inthe evening. The animals were provided with fresh filtered tap water adlibitum via an automatic watering system. During quarantine and thestudy, the animals were housed in an area where environmental controlswere set to maintain a temperature of 61 to 81° F. and a relativehumidity of 30 to 70%. Periodically the humidity was slightly out ofrange. A 12-hour light/dark cycle was employed and the room underwent aminimum of ten fresh air changes/hour. One of the seven animalsimplanted with the test articles (Group 3) was electively euthanized forhistopathological evaluation at the one month time point (Day 28). Allother animals were euthanized during the course of the study. Animalswere fasted for at least 12-24 hours prior to biopsy and implantationprocedures. Water was not withheld.

Preoperative Procedures.

Anesthesia and Analgesia. For biopsy and definitive surgery, animalswere sedated via intramuscular (IM) injection of a cocktail containing20 mg/kg ketamine, 2 mg/kg xylazine, and 0.040-mg/kg atropine. Eachanimal was then intubated and received inhalant isoflurane at 2.5%-4%for induction and 0.5-2.5% for maintenance of anesthesia, deliveredthrough either a volume-regulated respirator or rebreathing apparatus.Lactated Ringer's solution was administered at 10 ml/kg/hr for theduration of the surgical procedure. For technical procedures, animalswere sedated with the same intramuscular cocktail injection as describedabove. Ten ml of 10-mg/ml Propofol was also used in some animals at thediscretion of the attending veterinarian. For postoperative analgesia,Fentanyl patches (75 μg/hr) were applied to provide continuous painrelief. Alternatively, Rimadyl (50 mg) or Buprenex (0.05 mg/kg) wereadministered as needed.

Antibiotic Therapy. Broad-spectrum antibiotic therapy (approximately 5mg/kg Naxcel) was administered to all animals at the biopsy andimplantation surgeries. The treatment continued for up to 9 days and asneeded post surgery.

Surgical Preparation. For both biopsy and implantation surgery, the hairover the entire abdominal region was clipped. The animal was thenpositioned in dorsal recumbency. The operative area was cleaned withthree alternating scrubs of povidone-iodine solution and 70% alcohol;once the alternating scrubs are complete, a final application ofpovidone-iodine solution was applied and allowed to dry. The area(s) wasthen draped for aseptic surgery.

Surgical Procedures.

Biopsy/Tissue Collection. For all animals (Groups 1-3), biopsies ofadipose tissue, as well as venous blood, were obtained 10-11 weeks priorto Day 0 (implantation procedure). For tissue biopsy procedures, amidline incision was made in the abdomen beginning immediately caudal tothe umbilicus. Adipose biopsies of 21-34 grams of soft, pliablesubcutaneous adipose tissue (without connective tissue) were collectedaseptically from this midline access point. Collected tissue sampleswere individually and aseptically transferred to containers with tissueculture media (supplied by the Sponsor), then packaged in a bio-shipper(supplied by the Sponsor) and shipped overnight to the Sponsor forprocessing. The abdominal incision was closed in layers with absorbablesuture material of an appropriate size. The skin was closed in asubcuticular fashion, again using an appropriate size of absorbablesuture material. Approximately six 10-ml aliquots of venous blood werecollected in heparinized vacutainers, packaged in an ice-pack cooledcontainer (˜8° C.) and shipped overnight to the Sponsor for processing.

Cannulation procedure. At 12-18 days prior to test article implantation,an indwelling catheter was placed within the jugular vein of each animalto facilitate blood collection. The area surrounding the right jugularvein was shaved and prepared as described above. All animals werecannulated with a sterile 5.5-mm ID silicon catheter, which was insertedinto the right external jugular vein and secured by suture to preventmovement. An extra-large DaVINCI port was attached and implanted in asubcutaneous pocket.

Test Device Implantation. A midline abdominal incision was made 5 cmcranial to the umbilicus and extended approximately 15 cm caudally. Theperitoneum was identified and then carefully separated from theabdominal wall starting from the linea alba at midline and bilaterallytowards the right and left side of the abdominal wall. Care was taken toensure the tissue remained intact and vascularized. The urinary bladderwas then exposed and carefully emptied of urine, ensuring no urineentered into the abdominal cavity. The arteries and veins supplying thebladder were identified and ligated. The ureters were identified,stented (two 14-cm 7Fr DaVINCI non-absorbable ureter stents, inserted inascending fashion) and carefully transected from the bladder. Theurethra was over-sewn as it was transected. The bladder was thenremoved. The left ureter was carefully freed from the surroundingretroperitoneal fascia extending cranially until there was enoughmobility to reach the right side of test article. The right ureter wasdissected free to reach the other side of the test article. The ureterswere sutured on to the test article with 3-0 Vicryl in a simplecontinuous pattern. The peritoneum was transected on the left side andused to wrap the implants towards the right of midline portion whichprovided the vascular source and a watertight urine channel, and formeda tubular connection (atrium) between the caudal end of the implant(located in the intra-abdominal cavity) and the skin. The implant'scaudal end terminated in the peritoneal atrium approximately 5 to 7 cmaway from the skin stoma. The peritoneum was sutured with 3-0 Vicryl.The atrium was extended using the cranial peritoneal wrap whichtraversed the abdominal wall and exited the skin near the xiphoid (offmidline, right side). The externalized peritoneum was sutured to theskin to form a peritoneum-cutaneous junction and peritoneal-lined stomalumen. Surgical adhesive was then placed along the suture line where theperitoneum exited the body wall. The suture strands that were connectedto the ureteral stents were exteriorized through the stoma for futureremoval. The abdominal incision was closed with non-absorbable Prolenesuture. The skin was closed in a routine fashion. A Foley catheter wasinserted into the stoma to allow urine passage during stoma healing. Thesame surgical procedure was used for all animals.

Monitoring Procedures. Vital signs (oxygen rate, oxygen [O2] saturation,pulse rate, respiration, and body temperature) were monitored atintervals of approximately 20 minutes throughout the procedure.

Post Operative Procedures

Recovery. All animals were recovered after each surgical procedurewithin their own cages under normal environmental conditions.

Operative Drug Therapy. In addition to the antibiotic and analgesictherapies describes previously, Flomax was also used at the discretionof the facility veterinarian following surgeries and on as needed basisin order to maintain general good health during the survival period.

Stoma Maintenance. For two weeks post implantation or until the incisionsite was healed, the surgical area was evaluated for any signs ofdehiscence, abnormal discharge, odor, irritation or any abnormalities.The stoma area and surrounding tissue was cleaned twice daily, and thestoma catheter was observed for urine drainage. When not dripping, thecatheter was flushed with sterile saline to confirm patency. If thestoma became clogged, the flocculents and clogging materials wereremoved by forceps following the saline flush. If these efforts did notrestore the free flow of urine, a new catheter was installed and securedin place with prolene sutures. A stoma port was also installed andsecured with 2-0 prolene suture.

Stent Removal. At various time points from 2-4 weeks post-implantationsurgery the animals were anesthetized as described above and theureteral stents were removed.

Jugular Port. The jugular port catheter was flushed with injectablesaline and locked with Heparin (100 U/mL, ˜2-3 mL) weekly until Week 4and at each subsequent use to assure patency.

Imaging.

Ultrasonography. Ultrasound imaging of the conduit and kidneys wasperformed at Weeks 2, 4, 8 and prior to necropsy

Cystoscopy. Cystoscopy was performed at week 4 by inserting a bladderscope (flexible optical fiber with lens like a telescope or microscope)to view the inner surfaces of the conduit and to remove ureteral stentsas discussed above. Animals were anesthetized for this procedure.

Observations and Health Assessments. From the time of receipt untileuthanasia, the animals were observed twice daily for abnormalities ofappearance and behavior that might indicate adverse effects on health.At each check (performed approximately 8 hours apart), it was confirmedwhether all the animals had eaten and whether there was evidence ofurine and fecal output in each pen. Physical signs such as lethargy,emaciation, abnormal vocalization, missing anatomy, and laceration, aswell as abnormal behavioral signs, were noted, if present. Animals werenot removed from their pen during these daily assessments. Any abnormalsigns were documented.

Body weights. Body weights were recorded at baseline, weekly and priorto necropsy.

Clinical Pathology.

Blood Collection. Blood samples for analysis of hematology (CBC),coagulation, and serum chemistry parameters and were collected atscheduled time points (baseline, Week 1, 2, 3, 4, and 8, and prior tonecropsy) via the indwelling port in the jugular vein.

Hematology. Hematology samples were collected in 2.0 ml EDTA tubes andstored refrigerated on or wet ice (2-8° C.) prior to shipment (on wetice) to Idexx. Shipment was timed to facilitate analysis within 24 hoursof collection, as specified in the protocol. Samples were evaluated forthe hematology parameters: Total leukocyte count (WBC); Erythrocytecount (RBC); Hemoglobin concentration (HGB); Hematocrit value (HCT) 1;Mean corpuscular volume (MCV); Mean corpuscular hemoglobin (MCH) 1; Meancorpuscular hemoglobin concentration (MCHC) 1; Platelet count (PLT);Relative reticulocyte count (RTC), wherein 1=Calculated values.

Coagulation. Coagulation samples were collected in 1.8-ml sodium citratetubes (0.2 mL of 3.8% sodium citrate) and stored on wet ice until beingcentrifuged at 8,000 RPM for 10-15 minutes. The plasma was removed anddivided between two labelled vials, then frozen at −70° C. One vial waspackaged on dry ice and sent to Idexx for analysis, and the remainingvial was stored as a reserve until the conclusion of the study. Sampleswere evaluated for the following parameters: Prothrombin time (PT);Activated partial thromboplastin time (APTT); Fibrinogen (FIB).

Serum Chemistry. Blood samples for serum chemistry analysis werecollected in approximately 4.0-ml serum separation tubes. The bloodsamples were centrifuged at 10,000 RPM for 10-15 minutes and the serumwas extracted using sterile technique. Serum was divided between twolabeled vials and frozen at −70° C. One vial was packaged on dry ice andsent to Idexx for analysis, and the remaining vial was stored as areserve until the conclusion of the study. Samples were evaluated forthe following serum chemistry parameters: Glucose (GLU); Urea nitrogen(BUN); Creatinine (CRE); Total protein (TPR); Albumin (ALB); Globulin(GLOB) 1; Albumin/Globulin ratio (A/G) 1; Calcium (CAL); Phosphorus(PHOS); Sodium (NA); Potassium (K); Chloride (CL); Total cholesterol(CHOL); Total bilirubin (TBIL); Triglycerides (TRG); Alanineaminotransferase (ALT); Aspartate aminotransferase (AST); Alkalinephosphatase (ALK); Gamma glutamyltransferase (GGT); where 1=Calculatedvalues.

Blood Gases. Arterial blood gas samples were collected in a syringe(˜1.0 mL), and the blood was placed into a CG8+ i-STAT cartridge forin-house analysis. Samples were evaluated for the following blood gasparameters: Sodium (Na) (mmol/L) PCO2 (mm Hg); Potassium (K) (mmol/L)PO2 (mm Hg); Ionized Calcium (iCa) (mmol/L) TCO2 (mmol/L); Glucose (Glu)(mg/dL) HCO3 (mmol/L); Hematocrit (Hct) (%) BEecf (mmol/L); pH; andSO2(%).

Urine Collection. Urine samples were collected at baseline andpre-necropsy. Approximately 1.0 mL and 3.0 mL samples were collected insterile containers for qualitative and quantitative analysis,respectively. The qualitative analyses were done at the time ofcollection using Multistix® 10 SG Test Strips. The samples forquantitative comprehensive urinalysis were refrigerated and shipped toIDEXX Laboratories (North Grafton, Mass.) within 24 hours of collection.Samples were evaluated for the following qualitative urinalysisparameters: Glucose; Bilirubin; Blood; pH; Protein; Ketones;Urobilinogen; Specific Gravity; Nitrites; and Leukocytes, and for thefollowing quantitative urinalysis: Bacterial culture; Total bacteria;glucose; and total protein.

Anatomic Pathology

Moribund and Early Death Animals. Animals unlikely to survive until thenext scheduled observation (i.e., moribund animals) were weighed,euthanized, and necropsied. Animals were subjected to a limitednecropsy, aimed at determining a cause of declining health or death.Tissue collection from these specimens was limited to the urogenitalsystem. Necropsy occurred on the day of death, unless otherwise noted.An attempt was made to collect terminal samples for clinical pathologyfrom animals that were euthanized moribund.

Physical Examination. The animals were evaluated by the Test Facilityveterinarian prior to euthanasia. The general condition of the animalwas recorded.

Euthanasia. Animals were injected with sodium pentobarbital (150 mg/kg,IV) to cause euthanasia.

Necropsy. For the animal that was euthanized on schedule (84±5 days),the necropsy focused specifically on the kidneys, conduit, ureters,uretero-vesical junctions, mid-conduit, conduit-skin junction, and lymphnodes (lumbar and mesenteric).

Tissue Collection. At euthanasia and necropsy, all tissues surroundingthe neo-conduit implant were examined grossly and photographed in situ.The kidneys, the neo-conduit with attached ureters, and lymph nodes wereharvested. The kidneys were dissected and photographed. The neoconduitwith attached ureters was evaluated and pressure perfused with 10%normal buffered saline (NBF). All collected tissues were stored in 10%normal buffered saline (NBF) solution for histological processing.

Histology and Histopathology. Urinary organs were trimmed, examined,embedded in paraffin, and sectioned. Specific details regarding thesample sectioning scheme are included in the Pathology Report (below).Slides were stained with hematoxylin and eosin (H & E) and Masson'sTrichrome.

Results

Biopsy. Adipose tissue biopsies (21-34 grams) and venous blood (6×10 mLaliquots in heparinized tubes) were collected as outlined in theprotocol. Individual weights of adipose biopsy samples are presented inAppendix 1 (EE=Elective Euthanasia; PCV-2=Porcine Circovirus-2;SMC=Smooth Muscle Cells; S=Survivor; UD=Unscheduled Death due to poorclinical condition; X=observed).

Implantation (Surgical Methodology). All animals were recovered fromimplantation surgery uneventfully and the stoma was visualized to bedraining urine. The animal model was considered appropriate forevaluating the surgical application, post-surgical care andfunctionality of the Neo-Urinary Conduit.

FIG. 26 shows the ureteral stents used for the study. FIG. 27 shows theneo-conduit construct. FIG. 28 shows the neo-conduit construct attachedto the ureters. FIG. 29 shows the inflow end of the construct attachedto the ureters and the outflow end directed towards the surgicallycreated stoma. FIG. 30 shows the stoma and catheter for urine drainage.

Mortality. One of 7 animals survived until scheduled sacrifice (animal 4of Group 2, 83 days). Six of 7 animals were sacrificed unscheduled:animal 5 of Group 3 was electively euthanized 28 days post-implantationfor histopathological analysis and 5 animals were euthanized for poorclinical condition between 38 and 63 days post-implantation. (animal 1of Group 1, animals 2 and 3 of Group 2, and animals 6 and 7 of Group 3).These unscheduled deaths occurred in all treatment groups and wereattributed to viral infection and/or obstruction-related pathology withdamage to the upper urinary tract.

Final disposition and mortality findings for each of the 7 animals areshown in the table 3.2 below (EE=Elective Euthanasia; PCV-2=PorcineCircovirus-2; SMC=Smooth Muscle Cells; S=Survivor; UD=Unscheduled Deathdue to poor clinical condition; X=Observed)

TABLE 3.2 Cell Adipose Implant Seeding tissue Blood Date Animal SMCDensity collected collected (days after Stent Days on Mortality FindingsNumber Group Source (×10⁷) (grams) (ml) biopsy) Removal StudyDisposition PCV-2 Obstruction 1 1 Scaffold 0 24 60 74 Week 2 (1) 47 UD XOnly and week 4 (1) 2 2 Blood 2.5 21 60 71 Week 2 38 UD X X 3 2 Blood 230 60 72 Necropsy 40 UD X 4 2 Blood 2.5 25 60 71 Week 2 83 S X 5 3Adipose 2 34 60 73 Week 3 28 EE 6 3 Adipose 2 22 60 73 Week 2 (1) 39 UDX and week 3 (1) 7 3 Adipose 2.5 21 60 72 Week 4 63 UD X

Porcine Circovirus-2 (PCV-2) Infections. Three of 7 animals developedone or more lesions consistent with distinct pathological featuresascribed to porcine dermatitis and nephropathy syndrome (PDNS),associated with PCV-2 infection. Animals were classified asPCV-2-infected if the following were observed: 1) Clinically-observedpurple skin discoloration; 2) Microscopic vasculitis orvasculitis/perivasculitis affecting the kidney; 3) Kidney findings oftubular necrosis/fluid/casts/glomerulonephritis, or viral inclusions oftubular epithelial cells. By these criteria, porcine PCV-2 infection wasconfirmed in 3/7 animals. Two PCV-2 infected animals were unschedulednecropsy animals. These included animal 2 of Group 2 euthanized on day38 and animal 7 of Group 3 euthanized on day 63. The third animalidentified with PCV-2 infection (animal 4 of Group 2) survived toscheduled sacrifice (83 days).

Obstruction. Obstruction of urine flow through the conduit and stomacontributed to the morbidity in 4/6 unscheduled death animals. Theseincluded animal 1 of Group 1 euthanized on day 47; animals 2 and 3 ofGroup 2 euthanized on day 38 and day 40; and animal 6 of Group 3euthanized on day 39. Obstruction appeared to have been facilitated bythe placement of the test article in the ventral portion of theabdominal cavity of the quadruped where the weight of the overlyingabdominal organs could lead to conduit closure, adhesion and fistulaformation, and renal complications (e.g., dilation, inflammation, and/orinfection of ureters or kidney). Obstruction was exacerbated by the useof peritoneum to form the atrium, causing partial or full urinaryobstruction with subsequent detritus build-up and bacterial infection.Surgical placement of the test article was the same in all studyanimals; therefore, obstruction-related complications had a similarpathobiological mechanism (i.e. abdominal viscera resting on the conduitbecause of the quadruped anatomy) in all groups.

Clinical Heath Observations and Post-Surgical Care

Individual clinical health observations and post-surgical care for allanimals were observed (data not shown). To distinguish between theprotocol objective of establishing the post-surgical care followingimplantation of the test article and the ancillary 3-month pilotevaluation of the test article itself, the data below is broken into 2categories: observations <30 days and observations >30 days.

Clinical Heath Observations and Post-Surgical Care <30 Days. Followingtest article implantation surgery for all 7 animals, an indwelling Foleycatheter was placed through the stoma to facilitate urine drainage. All7 animals required weekly and on as needed basis maintenance of thiscatheter during the first 30 days postimplantation (e.g., Foley catheterreplacement [if dislodged], stoma flushing, and cleaning of the expecteddebris from anticipated scaffold biodegradation). This was performed aminimum of 8 and a maximum of 15 times overall during the first 30 days.Clinically, 7/7 animals experienced a loss of appetite (anorexia) orappeared thin during the first 30 days post-implantation. This included1/1 animals in Group 1 (animal 1); 2/3 animals in Group 2 (animals 2 and3) and 3/3 animals in Group 3 (5, 6, and 7). Seven of 7 animals appearedlethargic during the first 30 days post-implantation. This included 1/1animals in Group 1 (animal 1); 3/3 animals in Group 2 (animals 2, 3, and4) and 2/3 animals in Group 3 (animals 5 and 7). These clinicalobservations are not uncommon following urinary tract surgery in pigs.The following table 3.3 provides clinical health observations andpost-surgical care by test group (<30 Days Post implantation).

TABLE 3.3 Group 1 Group 2 Group 3 Clinical Observations N = 1 N = 3 N =3 Total N = 7 Stoma Maintenance 1/1 3/3 3/3 7/7 Anorexia (Not Eating,Thin) 1/1 3/3 3/3 7/7 Lethargy 1/1 3/3 3/3 7/7

Clinical Heath Observations and Post-Surgical Care >30 Days. Six animalswere on study >30 days (animal 5 of Group 3 was electively euthanized onday 28). One important clinical observation was intermittent obstructionof the outflow of urine. Since biodegradation of the scaffold occursduring the first 30 days post-implantation, obstruction appeared to havebeen facilitated by the placement of the test article in the ventralportion of the abdominal cavity of the quadruped where the weight of theoverlying abdominal organs could lead to conduit closure, adhesion andfistula formation, and renal complications (e.g., dilation,inflammation, and/or infection of ureters or kidney). Obstruction wasexacerbated by the use of peritoneum to form the atrium, causing partialor full urinary obstruction with subsequent detritus build-up andbacterial infection. Therefore, stoma maintenance was continued for thesix surviving animals from day 31 to necropsy. This was performed aminimum of 4 and a maximum of 13 times overall from day 31 to necropsy.Two of 6 animals presented with anorexia after day 30 (animal 1 of Group1 and animal 7 of Group 3). Three of 6 animals appeared lethargic afterday 30. Two animals presenting with lethargy in the first 30 dayscontinued to present with lethargy (animal 1 of Group 1 and animal 4 ofGroup 2) after day 30. The one animal that did not present with lethargyin the first 30 days presented with lethargy after day 30 (animal 6 ofGroup 3). The following table 3.4 shows clinical health observations andpost-surgical care by group (>30 Days post implantation).

TABLE 3.4 Group 1 Group 2 Group 3 Clinical Observations N = 1 N = 3 N =2* Total N = 6 Stoma port Maintenance 1/1 3/3 2/2 6/6 Anorexia (NotEating, Thin) 1/1 0/3 1/2 2/6 Lethargy 1/1 1/3 1/2 3/6 *animal 5 ofGroup 3 was electively euthanized at day 28.

Stent Removal. Stents were removed from anesthetized animals between 2and 4 weeks postimplantation by stent tether or visualization byultrasound and retrieval with cystoscope. The stents in one animal(animal 3 of Group 2) were not able to be visualized and removed andremained in place until necropsy. Individual ureteral stent removal datawas collected (data not shown).

Stoma port/Catheter. Individual data was collected (data not shown). Aprogression of changes was made during the study to optimize stomamanagement. Initially a Foley catheter was utilized. This was replacedwith a TRACOE® stoma ports. When this proved to be impractical due tofrequent device dislodgment by the animal, an 8Fr Foley catheter wasused to aid urine drainage. This was later replaced by a modifiedextension set tubing with bigger diameter to reduce clogging.Maintenance or replacement was done weekly and on an as needed basis.

Body Weights. All animals lost weight post-implantation. Body weightsfor all animals fluctuated during the course of the study. One animal(animal 4 of Group 2) survived until scheduled sacrifice, and remainedsteady or gained weight from week 2 until necropsy. Individual bodyweight data appears in the table 3.5 below.

TABLE 3.5 Animal No. Implant Type Baseline Wk 1 Wk 2 Wk 2 Wk 3 Wk 4 Wk 5Wk 6 Wk 7 Wk 8 Wk 9 Wk 12 1 Scaffold only 17.8 16.3 16.4 15.2 15.4 16.1NP 15.7 14.95 NA 2 Blood derived 20.1 19.1 NP 17.4 16.5 17.9 15.9 NA SMC3 Blood derived 17.4 15.9 16.3 16.3 16.6 17.25 NP 18.1 NP 19.6 19.8 19SMC 4 Blood derived 17.4 15.5 16.1 NP 17.1 19.3 18.1 NA SMC 5Adipose-derived 18.6 16.9 15.7 14.4 15.5 14.9 NP NP NP 16.65 13.4 NA SMC6 Adipose-derived 17.1 16.6 16.2 16.1 15.4 16.6 NP 17.2 NA SMC 7Adipose-derived 16.9 15.1 14.3 13.7 12.5 11.3 NA SMC

Clinical Pathology. Clinical pathology data for individual animals wascollected.

Hematology. Hematology data was collected (data not shown).Postoperative blood collection for hematology revealed the developmentof leukocytosis for all groups. Leukocyte counts for the all groupsfluctuated; however, leukocyte counts for the scaffold-only animal(Group 1) had the highest value at necropsy. Red blood cell counts(RBCs) fluctuated for all groups but remained within the reference range(8-10 MILL/uL) throughout the study. Hematocrit (%) fluctuated over thecourse of the study for all groups.

Serum Chemistry. Serum chemistry data was collected (data not shown).Overall, BUN, creatinine, total protein and potassium were elevated atnecropsy for all animals. Albumin was generally decreased by necropsy,and potassium and sodium fluctuated. Although changes were apparent inall groups, the scaffold only animal (Group 1) appeared to have the mostsignificant changes by necropsy.

Coagulation. Coagulation data was collected (data not shown). Theactivated partial thromboplastin time increased but then returned tonear baseline by necropsy for all animals except animals 5 and 7, whichremained elevated. Fibrinogen was elevated for all groups at necropsy,and was highest in the scaffold only animal.

Urinalysis. Urinalysis data was collected (data not shown). Urinalysisdata showed increases in protein, blood presence, white blood cell countand bacteria count for all groups between baseline and necropsy.

Imaging. Ultrasound data is presented in the following table 3.6.

TABLE 3.6 Week 2 Week 4 Right Left Conduit Right Left Conduit AnimalGroup Kidney Kidney Wall Kidney Kidney Wall No. Type L W L W Thickness LW L W Thickness 1 Scaffold 5.67 3.04 5.49 2.79 0.236 4.55 2.71 7.09 4.360.132 only 2 Blood 4.52 3.33 5.55 3.96 0.139 9.45 3.54 8.08 3.59 0.16 4derived 5 2.91 6.23 3.94 0.272 10.5 4.31 5.2 3.49 0.17 3 4.5 2.57 5.873.2 0.172 6.47 3.7 5.39 3.18 0.176 6 Adipose 4.33 3.07 5.67 3.58 0.17111.7 5.37 7.09 4.47 0.334 5 derived 5.18 3.26 5.65 2.77 0.246 4.1 2.15.08 2.76 0.147 7 4.82 2.62 3.57 2.14 0.233 10.2 3.5 7.09 3 0.403 Week 8Conduit Week 12 Right Left Wall Right Left Conduit Animal Group KidneyKidney Thickness Kidney Kidney Wall No. Type L W L W (cm) L W L WThickness 1 Scaffold NA NA only 2 Blood NA NA 4 derived 8.05 3.32 7.054.85 0.22 7.16 4.19 8.35 3.59 0.242 3 NA NA 6 Adipose NA NA 5 derived NANA 7 7.51 4.4 7.09 3.31 0.211 NA

FIGS. 31-43 show representative cystoscopy images. FIG. 31 shows acytoscopy image of animal 1 of Group 1 at week 4 (cell-free scaffoldimplanted). FIG. 32 shows the same animal 2 days before necropsy at the6 week check-up.

FIG. 33 shows the cytoscopy image of animal 2 of Group 2 pre-necropsy(scaffold seeded with blood-derived SMCs).

FIGS. 34-36 shows the cytoscopy image of animal 3 of Group 2 at week 4,week 5, and pre-necropsy (scaffold seeded with blood-derived SMCs).

FIG. 37 shows the cytoscopy image of animal 4 of Group 2 pre-necropsy(scaffold seeded with blood-derived SMCs).

FIG. 38 shows the cytoscopy image of animal 5 of Group 3 at week 3(scaffold seeded with adipose-derived SMCs). The image shows mucosa thatis covered by white-tan amorphous granular casts of scaffold debris.FIG. 39 shows an image of the same animal at week 3. The pinkish focusnear or at the ureteral anastomoses indicates where the epithelium cameoff with removal of the stents leaving a pink, vascularized granulationbed.

FIGS. 40-42 shows the cytoscopy image of animal 6 of Group 3 at week 3,week 4, and pre-necropsy (scaffold seeded with adipose-derived SMCs).

FIG. 43 shows the cytoscopy image of animal 7 of Group 3 at week 4(scaffold seeded with adipose-derived SMCs). Two stents (blue material)were initially seen under ultrasound imaging and were later removed withcytoscopic guidance.

Ultrasound. Individual and group ultrasonography data are presented inthe table above. Ultrasonography on the kidneys over the course of thestudy showed increases in surface area (length×width) indicating kidneychanges (hydronephrosis) in all treatment groups. The following table3.7 provides the kidney surface area (L×W, cm2).

TABLE 3.7 Group Animal Week No. Group Type No. Week 2 Week 4 Week 8 12Left Kidney 1 Scaffold only 1 15.32 30.91 NA NA 2 Blood derived 2 21.9829.01 NA NA 2 Blood derived 3 24.55 18.15 34.19 29.98 2 Blood derived 418.78 17.14 NA NA 3 Adipose derived 5 20.30 31.69 NA NA 3 Adiposederived 6 15.65 14.02 NA NA 3 Adipose derived 7 7.64 21.27 23.47 NARight Kidney 1 Scaffold only 1 17.24 12.33 NA NA 2 Blood derived 2 15.0533.45 NA NA 2 Blood derived 3 14.55 45.26 26.73 30.00 2 Blood derived 411.57 23.94 NA NA 3 Adipose derived 5 13.29 62.83 NA NA 3 Adiposederived 6 16.89 8.61 NA NA 3 Adipose derived 7 12.63 35.70 33.04 NA

Ultrasonography on the implant wall thickness showed a fluctuation inthickness for all Groups. The following table 3.8 illustrates this.

TABLE 3.8 Conduit Wall Thickness (cm) Group Animal Week No. Group TypeNo. Week 2 Week 4 Week 8 12 1 Scaffold only 1 0.236 0.132 NA NA 2 Bloodderived 2 0.139 0.160 NA NA 2 Blood derived 3 0.272 0.170 0.220 0.242 2Blood derived 4 0.172 0.176 NA NA 3 Adipose derived 5 0.171 0.334 NA NA3 Adipose derived 6 0.246 0.147 NA NA 3 Adipose derived 7 0.233 0.4030.211 NA

The pathology report appears in Example 4 below.

Evidence of Porcine Circovirus Type-2 (PCV-2) infection was observed in3/7 animals. These included animal 2 of Group 2 euthanized on day 38 andanimal 7 of Group 3 euthanized on day 63. The third animal identifiedwith PCV-2 infection, animal 4 of Group 2, survived to scheduledsacrifice (83 days). Obstruction of urine flow through the conduit andstoma contributed to the morbidity in 4/6 unscheduled death animals.These included animal 1 of Group 1, euthanized on day 47; animals 2 and3 of Group 2 euthanized on days 38 and 40; and animal 6 of Group 3euthanized on day 39. Ventral abdomen positioning of surgicallyimplanted test article contributed to physical obstruction of urine flowin the quadruped animal model where the weight of the overlyingabdominal organs contributed to conduit closure, adhesion and fistulaformation, and secondary upper urinary tract renal complications (e.g.,dilation, inflammation, and/or infection of ureters or kidney). Inaddition, the urine flow obstruction was exacerbated by the use ofperitoneum to form the atrium, causing partial or full urinaryobstruction with subsequent detritus build-up and bacterial infection.Surgical placement of the test article was the same in all studyanimals; therefore, obstruction-related complications had a similarpathobiological mechanism in all groups. Regeneration of urinary-liketissue was evident as early as day 28, with presence of urothelium,lamina propria and smooth muscle bundles at the ureter-conduit junction(UCJ) in an electively euthanized animal (animal 5 of Group 3,adipose-derived SMC). The regenerative process at the ureteral end ofthe implant resulted in urinary-like tissue formation that wascomparable among animals receiving a construct implant (Groups 2 and 3).The extent of urinary-like tissue regeneration in the construct groups(Groups 2 and 3) was influenced by duration of animal survivalpost-implantation. The one animal surviving to scheduled sacrifice(animal 4 of Group 2 at day 83) had urothelium and smooth muscle presentin the UCJ, cranial and mid portions of the conduit in spite of adetected viral infection. However, the peritoneum-only atrium wasinsufficient to support urinary-like tissue regeneration and the tissueformed in the atrium had a wall comprised of fibrous connective tissuewithout urothelial mucosal lining The point of transition from conduitto atrium varied between animals because the caudal end of the implantfloated freely within the peritoneal wrapping making the transition fromconduit to atrium difficult to define at necropsy. The typicalcomposition of (presumed) caudal conduit was organized collagen withassociated fibroblasts and/or myofibroblasts. Peritoneum atrium appearsto be insufficient for urinary-like tissue regeneration, but does serveas a source of vascularization to NUC implants.

CONCLUSIONS. The swine animal model proved appropriate for evaluatingthe surgical application of the Neo-Urinary Conduit in this studybecause all animals recovered from surgery and urinary diversion wasachieved. In addition, the swine model was appropriate for evaluatingpost-operative care of urinary flow obstruction and its impact to theupper urinary tract. Finally, the swine model was appropriate forevaluating the ability of the test articles to regenerate urinary-liketissue in an environment complicated by detritus accumulation andbacterial colonization, viral infection, and enteric adhesions andfistulas.

The surgical methodology was determined to be successful althoughanatomical placement of the urinary diversion on the ventral abdominalfloor of a quadrupedal animal resulted in partial obstruction of urineoutflow. The animal model was considered appropriate for evaluating thesurgical application, postsurgical care and functionality of theNeo-Urinary Conduit.

Post-surgical findings during the first 30 days following implantsurgery revealed findings that were not considered uncommon followingurinary diversion surgery in the pig.

Although several confounding factors occurred during the study (i.e.surgical placement on the ventral abdominal floor, use of peritonealatrium and viral infection), regeneration of urinary-like tissue wasevident as early as day 28, with presence of urothelium, lamina propriaand smooth muscle bundles at the ureterconduit junction in an electivelyeuthanized animal (animal 5 of Group 3, adipose-derived SMC).

The extent of urinary-like tissue regeneration in the construct groups(Groups 2 and 3) was influenced by duration of animal survivalpost-implantation. The one animal surviving to scheduled sacrifice(animal 4 of Group 2 at Day 83) had urothelium and smooth muscle presentin the UCJ, cranial and mid portions of the conduit in spite of adetected viral infection.

There were no apparent differences observed in the regenerative processwhen scaffolds were seeded with SMC derived from blood or adipose(Groups 2 and 3, respectively), suggesting equivalence between SMCsources in promoting regeneration.

The tissue formed from peritoneum in the atrium segment of the conduithad a wall comprised of fibrous connective tissue without urotheliallining.

Example 4 Pathology of Animals Following Implantation of Neo-UrinaryConduit Constructs

At the conclusion of the study described in Example 3, the anatomicpathology of the test animals was assessed.

Tissue Collection. The abdominal cavity was opened and the conduit,i.e., the outcome of implanting a construct or scaffold only testarticle, was visualized and digitally photographed in situ at the animalfacility. The conduit was removed en bloc with the kidneys and ureters.The ureters were detached from the conduit by transverse sectioning 3-4cm away from the anastomoses. Representative sections of the kidneys,ureters, lymph nodes, and any other lesions observed grossly wereobtained. All tissue samples were placed in 10% Neutral BufferedFormalin (NBF) for 24-48 hours prior to shipping to Vet Path Services,Inc. for histological processing and evaluation.

Histological Processing. After fixation, the conduit was openedlongitudinally (parallel with the outflow) and divided into dorsal andventral halves as illustrated in FIG. 44.

Three transverse sections were trimmed from each half (cranial, mid andcaudal sections were captured for the dorsal half and the ventral half).One section from each half was taken from the conduit-atrium junction.An additional section was taken at each of the two ureterconduitjunctions. One other slide was used to capture the stoma at the skinsurface and the adjacent canal through the abdominal wall. When the sizeof the conduit permitted, this scheme resulted in 11 slides. Sectionswere collected from each animal. In addition, the following tissue/organsections were obtained and submitted for histology: left kidney, rightkidney, left ureter, right ureter, lumbar lymph node, mesenteric lymphnodes inguinal lymph node and any gross lesions.

During trimming of tissues at VPS, digital photographs were taken forillustration purposes. Post fixation, tissues were processed routinelyto microslides and stained with hematoxylin and eosin (H&E) and Masson'strichrome. In addition, slides for the kidneys and 5 conduit sites werestained with Brown and Hopps (Gram) stain. Slides were evaluatedmicroscopically.

Where appropriate, microscopic observations for Individual Animal Datawere obtained and scored.

Results

Mortality. Animals survived 28-83 days. One of 7 animals survived untilscheduled sacrifice (an animal from Group 2, 83 days). Six of 7 animalswere sacrificed unscheduled: a Group 3 animal was electively euthanizedat 28 days post-implantation for histopathological analysis and 5animals were euthanized for poor clinical condition between 38 and 63days post-implantation (a Group 1 Animal; two Group 2 Animals; and twoGroup 3 Animals). These unscheduled deaths occurred in all treatmentgroups and were attributed to viral infection and/or obstruction-relatedpathology with damage to the upper urinary tract. Final disposition andmortality findings for each of the 7 animals are shown in Appendix 1,Animal Information. Disposition (mortality classification) is summarizedby treatment group in the following table 4.2 (Group 1=Scaffold-only,Group 2=Blood-derived SMC construct, Group 3=Adipose-derived SMCconstruct).

TABLE 4.2 Unscheduled and Scheduled Deaths by Group Group 1 Group 2Group 3 Total Mortality N = 1 N = 3 N = 3 N = 7 Unscheduled Necropsy(Euthanized) 1/1 2/3 3/3* 6/7 Scheduled Necropsy 0/1 1/3 0/3  1/7

Evidence of Porcine Circovirus Type 2 (PCV-2). Three of 7 animalsdeveloped one or more lesions consistent with distinct pathologicalfeatures ascribed to porcine dermatitis and nephropathy syndrome (PDNS),associated with PCV-2 infection. Animals were classified asPCV-2-infected if the following were observed: 1) Clinically-observedpurple skin discoloration; 2) Microscopic vasculitis orvasculitis/perivasculitis affecting the kidney; 3) Kidney findings oftubular necrosis/fluid/casts/glomerulonephritis, or viral inclusions oftubular epithelial cells.

By these criteria, porcine PCV-2 infection was confirmed in 3/7 animals.Two PCV-2 infected animals were unscheduled necropsy animals. Theseincluded one animal in Group 2 (animal 2) euthanized on day 38 andanother animal in Group 3 (animal 7) euthanized on day 63. The thirdanimal identified with PCV-2 infection, animal 4 of Group 2, survived toscheduled sacrifice (83 days).

Obstruction. Obstruction of urine flow through the conduit and stomacontributed to the morbidity in 4/6 unscheduled death animals. Theseincluded animal 1 of Group 1, euthanized on day 47; animals 2 and 3 ofGroup 2, euthanized on days 38 and 40; and animal 6 of Group 3,euthanized on day 39. Ventral abdomen positioning of surgicallyimplanted test article contributed to physical obstruction of urine flowin the quadruped animal model where the weight of the overlyingabdominal organs contributed to conduit closure, adhesion and fistulaformation, and secondary upper urinary tract renal complications (e.g.,dilation, inflammation, and/or infection of ureters or kidney). Inaddition, the urine flow obstruction was exacerbated by the use ofperitoneum to form the atrium, causing partial or full urinaryobstruction with subsequent detritus build-up and bacterial infection.Surgical placement of the test article was the same in all studyanimals; therefore, obstruction-related complications had a similarpathobiological mechanism in all groups.

Distribution of Unscheduled and Scheduled Deaths by Underlying Findings.A summary of underlying findings in the unscheduled and scheduled deathsis presented in the following table 4.3 (Group 1=Scaffold-only, Group2=Blood-derived SMC construct, and Group 3=Adipose-derived SMCconstruct).

TABLE 4.3 Group 1 Group 2 Group 3 Total Mortality Categories N = 1 N = 3N = 3 N = 7 Evidence of PCV-2 infection 0 2*{circumflex over ( )} 1 3(unscheduled and scheduled death) Obstruction 1 2{circumflex over ( )} 14 Elective Euthanasia 0 0 1 1 Total unscheduled deaths 1 2{circumflexover ( )} 3 6 Total scheduled deaths 0 1* 0 1 *animal 4 of Group 2survived until scheduled sacrifice (Day 83) {circumflex over ( )}animal2 of Group 2 showed evidence of PCV-2 and obstruction related findings

Surgical Care. A comprehensive list of macroscopic findings andmicroscopic correlates for all animals was obtained (data not shown).

Inter-group incidences for kidney, ureter and other findings are shownin the following tables. Table 4.6 below shows the intergroup incidence:kidney, ureter and other tissue findings (Number of animals with finding(of any severity)/Number of animals examined; and U=Unexamined).

TABLE 4.6 Group: Treatment: 2 3 1 Autologous Blood Autologous AdiposeScaffold Only SMC SMC Mean Days on Study: 47 54 43 Left KidneyHydronephrosis/chronic nephritis 1/1 1/3 1/3 Chronic nephritis (withouthydronephrosis) 0/1 1/3 0/3 Pyelonephritis 0/1 1/3 0/3 Chronic-activenephritis 0/1 1/3 3/3 Regeneration, tubular epithelium 1/1 2/3 2/3Tubular necrosis/fluid/casts/glomerulonephritis 0/1 0/3 1/3Vasculitis/perivascular inflammation 0/1 1/3 1/3 Viral inclusions 0/10/3 1/3 Inflammation, chronic-active, 0/1 0/3 1/3 capsule/peritoneumBacterial colonies, capsule/peritoneum 0/1 0/3 1/3 Right KidneyHydronephrosis/chronic nephritis 1/1 1/3 1/3 Chronic nephritis (withouthydronephrosis) 0/1 0/3 0/3 Pyelonephritis 0/1 1/3 0/3 Chronic-activenephritis 1/1 1/3 1/3 Regeneration, tubular epithelium 0/1 2/3 3/3Tubular necrosis/fluid/casts/glomerulonephritis 0/1 2/3 0/3Vasculitis/perivascular inflammation 0/1 1/3 0/3 Viral inclusions 0/10/3 0/3 Inflammation, chronic-active, 1/1 0/3 0/3 capsule/peritoneumBacterial colonies, capsule/peritoneum 1/1 0/3 0/3 Left UreterDilatation 0/1 1/3 1/3 Vacuolation, transitional epithelium 0/1 1/3 0/3Inflammation, subacute/chronic, peri-ureter 1/1 2/3 2/3 mesenteryVasculitis/necrosis, mesenteric blood vessel 0/1 1/3 2/3 Right UreterDilatation 1/1 1/3 2/3 Vacuolation, transitional epithelium 0/1 0/3 0/3Inflammation, subacute/chronic, peri-ureter 1/1 1/3 2/3 mesenteryVasculitis, mesenteric blood vessel 0/1 1/3 1/3 Lymph Node, Inguinal(Normal) 1/1 1/1 3/3 Lymph Node, Lumbar (Normal) U 1/2 2/3Histiocytosis, sinus U 1/2 1/3 Lymph Node, Mesenteric (Normal) U 2/2 2/3Infiltrate, neutrophils U 0/2 1/3 Adhesions and Fistulas Adhesionconduit to intestines (macroscopic 1/1 2/3 2/3 and microscopic) Adhesionconduit to intestines (microscopic 0/1 1/3 0/3 only) Adhesion conduit touterus (macroscopic) 0/1 1/3 0/3 Adhesion ureter to uterus/ovary(macroscopic) 0/1 1/3 1/3 Adhesion ureter to intestines (macroscopic)0/1 0/3 1/3 Fistula (macroscopic) & Fistula/neutrophil tract 0/1 2/3 0/3(microscopic) GROSS LESIONS: Skin, Inner Left Thigh & PerineumVasculitis, acute, necrotizing, dermis U 1/1 U Hemorrhage, dermis U 1/1U

Table 4.7 below shows the intergroup incidence: kidney, ureter and othertissue findings (Number of animals with finding (of any severity)/Numberof animals examined; and U=Unexamined) (non-PCV-2 animals).

TABLE 4.7 Group: Treatment: 2 3 1 Autologous Blood Autologous AdiposeScaffold Only SMC SMC Left Kidney Hydronephrosis/chronic nephritis 1/10/1 1/2 Chronic nephritis (without hydronephrosis) 0/1 0/1 0/2Pyelonephritis 0/1 0/1 0/2 Chronic-active nephritis 0/1 0/1 2/2Regeneration, tubular epithelium 1/1 0/1 1/2 Tubularnecrosis/fluid/casts/glomerulonephritis 0/1 0/1 0/2Vasculitis/perivascular inflammation 0/1 0/1 1/2 Viral inclusions 0/10/1 0/2 Inflammation, chronic-active, 0/1 0/1 1/2 capsule/peritoneumBacterial colonies, capsule/peritoneum 0/1 0/1 1/2 Right KidneyHydronephrosis/chronic nephritis 1/1 0/1 1/2 Chronic nephritis (withouthydronephrosis) 0/1 0/1 0/2 Pyelonephritis 0/1 0/1 0/2 Chronic-activenephritis 1/1 0/1 0/2 Regeneration, tubular epithelium 0/1 0/1 2/2Tubular necrosis/fluid/casts/glomerulonephritis 0/1 0/1 0/2Vasculitis/perivascular inflammation 0/1 0/1 0/2 Viral inclusions 0/10/1 0/2 Inflammation, chronic-active, 1/1 0/1 0/2 capsule/peritoneumBacterial colonies, capsule/peritoneum 1/1 0/1 0/2 Left UreterDilatation 0/1 0/1 1//2 Vacuolation, transitional epithelium 0/1 1/1 0/2Inflammation, subacute/chronic, peri-ureter 1/1 1/1 2/2 mesenteryVasculitis/necrosis, mesenteric blood vessel 0/1 0/1 2/2 Right UreterDilatation 1/1 0/1 2/2 Vacuolation, transitional epithelium 0/1 0/1 0/2Inflammation, subacute/chronic, peri-ureter 1/1 1/1 2/2 mesenteryVasculitis, mesenteric blood vessel 0/1 1/1 1/1 Lymph Node, Inguinal(Normal) 1/1 1/1 2/2 Lymph Node, Lumbar (Normal) U 1/1 2/2 Lymph Node,Mesenteric (Normal) U 1/1 2/2 Adhesions and Fistulas Adhesion conduit tointestines (macroscopic 1/1 1/1 1/2 and microscopic) Adhesion conduit tointestines (microscopic 0/1 0/1 0/2 only) Adhesion conduit to uterus(macroscopic) 0/1 0/1 0/2 Adhesion ureter to uterus/ovary (macroscopic)0/1 0/1 1/2 Adhesion ureter to intestines (macroscopic) 0/1 0/1 1/2Fistula (macroscopic) & Fistula/neutrophil tract 0/1 1/1 0/2(microscopic)

Conduits formed from implantation of a test article were variably sizedand shaped tubes located in the retro-peritoneal space of the ventralabdomen. The ureters entered at the cranial end of the conduit(ureter-conduit junction, UCJ, FIG. 44). Urine flow was directed throughthe peritoneal-wrapped implant and the atrium and emerged at the stoma.The cranial end of the conduit (ureteral attachment) frequently hadbilateral bulbous dilations, referred to as diverticula that wereconsidered to be part of the regenerative process and reflected theintermittent obstruction of the stoma and back pressure causing adilatation to develop in the regenerated conduit.

Adhesions and Fistulas. The ventral side of the conduit was adhered tothe fascia and skeletal muscle of the abdominal wall, and the dorsalside was covered with peritoneum. At necropsy, adhesions between theconduit or ureters and other abdominal organs (e.g., gastrointestinaltract, omentum or other abdominal organs) were observed. The lumen ofthe conduit was filled with detritus. Abdominal adhesions, includingadhesions among the various abdominal organs and adhesions between theconduit and abdominal organs, were present in 7/7 animals. The incidenceof adhesions and fistulas is summarized in the following Table 4.8 below(Group 1=Scaffold-only, Group 2=Blood-derived SMC construct, and Group3=Adipose-derived SMC construct)

TABLE 4.8 Group 1 Group 2 Group 3 Total Finding N = 1 N = 3 N = 3 N = 7Adhesion conduit to intestines 1/1 3/3 2/3 6/7 Adhesion conduit touterus 0/1 1/3 0/3 1/7 Adhesion ureter to uterus or ovary 0/1 1/3 2/33/7 Adhesion ureter to intestine 0/1 0/3 1/3 1/7 Fistula (conduit tointestines) 0/1 2/3 0/3 2/7

Six of 7 animals had adhesions between the conduit and intestine(macroscopically and microscopically in animal 1 of Group 1; animals 3and 4 of Group 2 and animals 6 and 7 of Group 3; microscopically inanimal 2 of Group 2). One of 7 animals also had macroscopic adhesionsbetween the conduit and uterus and ureter to uterus or ovary (animal 4of Group 2). One animal (animal 5 of Group 3) had an adhesion betweenthe ureter and uterus or ovary (macroscopically). One of 7 animals hadmacroscopic adhesions ureter to uterus or ovary and ureter to intestine(animal 6 of Group 3). Fistulas were observed macroscopically, andfistula/neutrophil tract microscopically, between the conduit andintestinal tract in 2/7 animals (animals 2 and 3 of Group 2). Fistulaswere observed macroscopically, and fistula/neutrophil tractmicroscopically, between the conduit and intestinal tract in 2/7 animals(animals 2 and 3 of Group 2).

Ureters and Kidneys. Thickened ureters observed macroscopically resultedfrom several underlying biological processes upon microscopicevaluation. Ureter dilatation (or hydroureter) was characterized by anexpanded lumen with normal ureteral wall structure. Ureters were alsosometimes thickened by subacute/chronic inflammation of the peri-uretermesentery, which occurred when the mesentery surrounding the ureter wasexpanded by collagen and fibroblasts, with occasional lymphocytes andmacrophages. Transitional cell vacuolation was characterized by round,clear vacuoles in the epithelium. This inflammation did not usuallyaffect the muscle tunics or urothelium of the ureter. Peri-ureterinflammation could be related to adhesion between ureter and intestinalor reproductive organs; but it could also occur without adhesion.Vasculitis, with or without necrosis of blood vessels, was observedwithin the areas of peri-ureter inflammation. This may have beenassociated with the PCV-2 viral infection in animals 2 of Group 2.Microscopically, hydronephrosis was characterized by a dilatation of therenal pelvis with thinning and chronic inflammation (fibrosis,lymphocytes, plasma cells and occasional macrophages) of the renalcortex. Hydronephrosis was considered to be the result of full orpartial obstruction in the lower urinary system (ureters, conduit oratrium/stoma). Chronic-active pyelonephritis, which was occasionallyassociated with hydronephrosis, was characterized by infiltration ofneutrophils and cellular debris into the renal pelvis, often spreadinginto the distal medulla. Pyelonephritis was the result of bacterialinfection of the lower urinary tract which ascended into the renalpelvis. Chronic nephritis (without hydronephrosis) was characterized byfibrosis with infiltration of inflammatory cells (lymphocytes,macrophages, plasma cells and occasionally neutrophils) in the renalcortex or medulla. The cortex of kidneys with chronic nephritis lookedsimilar to those in animals with hydronephrosis/chronic nephritis;however, in chronic nephritis the pelvis was not dilated. Chronic-activenephritis was similar in appearance to chronic nephritis, but withsignificant infiltration of neutrophils. Tubularnecrosis/fluid/casts/glomerulonephritis was a constellation of changescharacterized by neutrophils, lymphocytes and macrophages in glomeruli,necrosis of individual tubular epithelial cells, proteinaceous tubularcasts and/or hemorrhage in tubular lumens. Tubularnecrosis/fluid/casts/glomerulonephritis was observed in animals 2 and 4of Group 2 and animal 7 of Group 3. Also common in the kidneys of theanimals with tubular necrosis/fluid/casts/glomerulonephritis wasvasculitis/perivascular inflammation. Chronic-active inflammation of thecapsule/peritoneum was characterized by thickening of the capsule of thekidney by fibroblasts, collagen fibers and/or fibrin, neutrophils,lymphocytes and macrophages, and was indicative of peritonitis.

Microscopic Hydroureter and Hydronephrosis and/or Pyelonephritis.Hydroureter and hydronephrosis were observed in 4 of 7 animals. Theresults are shown in the following table 4.9 (Group 1=Scaffold-only,Group 2=Blood-derived SMC construct, and Group 3=Adipose-derived SMCconstruct).

TABLE 4.9 Incidence of Hydroureter, Hydronephrosis and/or PyelonephritisGroup 1 Group 2 Group 3 Finding N = 1 N = 3 N = 3 Total N = 7Hydroureter, Unilateral 1/1 0/3 1/3 2/7 Hydroureter, Bilateral 0/1 1/31/3 2/7 Hydronephrosis, Unilateral 0/1 0/3 2/3 2/7 Hydronephrosis,Bilateral 1/1 1/3 0/3 2/7 Pyelonephritis, Unilateral 0/1 0/3 0/3 0/7Pyelonephritis, Bilateral 0/1 1/3 0/3 1/7

Unilateral hydroureter (2/7 animals): 1 animal in Group 1 (animal 1) and1 animal in Group 3 (animal 6). Bilateral hydroureter (2/7 animals): 1animal in Group 2 (animal 2) and 1 animal in Group 3 (animal 5).Unilateral hydronephrosis (2/7 animals): 2 animals in Group 3 (animals 5and 6). Bilateral hydronephrosis (2/7 animals): 1 animal in Group 1(animal 1) and 1 animal in Group 2 (animal 2). Pyelonephritis(bilateral) was observed in 1 of 7 animals; Group 2 (animal 2). The oneanimal that survived to the scheduled sacrifice (Group 2 animal 4, Day83) did not have hydroureter, hydronephrosis or pyelonephritis.

Findings related to Regeneration of a Urinary Conduit. Detailed findingsof tissue components observed in each section harvested was collected(data not shown).

Inter-group incidences for regenerated conduit findings are shown in thefollowing tables. The table 4.17 below shows the NUC Summary of findingsby Group.

TABLE 4.17 Group: Treatment: 2 3 1 Autologous Blood Autologous AdiposeScaffold Only SMC SMC Mean Days on Study: 47 54 43 Incidence of SelectFindings by Conduit Location (# animals with finding/# animals examinedat that location) Presence of Urothelium Left Ureter Conduit Junction(Slide14) 1/1 1/3 3/3 Right Ureter Conduit Junction (Slide 16) 0/1 1/33/3 Cranial (Ureter) End of Conduit Body (Slides 6 or 0/1 1/3 0/3 10)Middle of Conduit Body (Slides 7 or 11) 0/1 1/3 0/3 Caudal ConduitBody/Atrium (Slides 8, 9, 12 or 13) 0/1 0/3 0/3 Atrium-Stoma-SkinJunction (Slide 18) 0/1 0/3 0/3 Presence of Squamous Epithelium inAtrium/Conduit Left Ureter Conduit Junction (Slide14) 0/1 0/3 0/3 RightUreter Conduit Junction (Slide 16) 0/1 0/3 0/3 Cranial (Ureter) End ofConduit Body (Slides 6 or 0/1 0/3 0/3 10) Middle of Conduit Body (Slides7 or 11) 0/1 0/3 0/3 Caudal Conduit Body/Atrium (Slides 8, 9, 12 or 13)0/1 1/3 0/3 Atrium-Stoma-Skin Junction (Slide 18) 0/1 2/3 1/3 SmoothMuscle Left Ureter Conduit Junction (Slide14) 1/1 0/3 1/3 Right UreterConduit Junction (Slide 16) 0/1 1/3 1/3 Cranial (Ureter) End of ConduitBody (Slides 6 or 0/1 1/3 0/3 10) Middle of Conduit Body (Slides 7 or11) 0/1 1/3 0/3 Caudal Conduit Body/Atrium (Slides 8, 9, 12 or 13) 0/10/3 0/3 Atrium-Stoma-Skin Junction (Slide 18) 0/1 0/3 0/3 Surface ofChronic-Active Inflammation/Detritus Left Ureter Conduit Junction(Slide14) 0/1 3/3 1/3 Right Ureter Conduit Junction (Slide 16) 1/1 2/31/3 Cranial (Ureter) End of Conduit Body (Slides 6 or 1/1 3/3 3/3 10)Middle of Conduit Body (Slides 7 or 11) 1/1 3/3 3/3 Caudal ConduitBody/Atrium (Slides 8, 9, 12 or 13) 1/1 3/3 3/3 Atrium-Stoma-SkinJunction (Slide 18) 1/1 3/3 2/3 Attenuation, Urothelium 1/1 0/3 2/3Hyperplasia, Urothelium 0/1 1/3 1/3 Vacuolation, Urothelium 0/1 0/3 1/3Hemorrhage, Conduit Wall 1/1 1/3 1/3 Scaffold Material, Conduit Wall 0/10/3 2/3 Increased Collagen, Scaffold Wall 1/1 0/3 0/3 Adhered GI Tract(P = present) 0/1 2/3 0/3 Fistula/neutrophil Tract to Intestines 0/1 2/30/3 Acanthosis, skin, stoma 0/1 2/3 1/3 Brown and Hopps Gram Stain Grampositive bacteria in detritus 1/1 3/3 3/3 Gram negative bacteria indetritus 1/1 3/3 3/3

Table 4.18 below shows the NUC Summary of findings by Group (Non-PCV-2Animals).

TABLE 4.18 Group: Treatment: 2 3 1 Autologous Blood Autologous AdiposeScaffold Only SMC SMC Mean Days on Study: 47 40 34 Incidence of SelectFindings by Conduit Location (# animals with finding/# animals examinedat that location) Presence of Urothelium Left Ureter Conduit Junction(Slide14) 1/1 0/1 2/2 Right Ureter Conduit Junction (Slide 16) 0/1 0/12/2 Cranial (Ureter) End of Conduit Body (Slides 6 or 10) 0/1 0/1 0/2Middle of Conduit Body (Slides 7 or 11) 0/1 0/1 0/2 Caudal ConduitBody/Atrium (Slides 8, 9, 12 or 13) 0/1 0/1 0/2 Atrium-Stoma-SkinJunction (Slide 18) 0/1 0/1 0/2 Presence of Squamous Epithelium inAtrium/Conduit Left Ureter Conduit Junction (Slide14) 0/1 0/1 0/2 RightUreter Conduit Junction (Slide 16) 0/1 0/1 0/2 Cranial (Ureter) End ofConduit Body (Slides 6 or 10) 0/1 0/1 0/2 Middle of Conduit Body (Slides7 or 11) 0/1 0/1 0/2 Caudal Conduit Body/Atrium (Slides 8, 9, 12 or 13)0/1 0/1 0/2 Atrium-Stoma-Skin Junction (Slide 18) 0/1 1/1 0/2 SmoothMuscle Left Ureter Conduit Junction (Slide14) 1/1 0/1 1/2 Right UreterConduit Junction (Slide 16) 0/1 0/1 1/2 Cranial (Ureter) End of ConduitBody (Slides 6 or 10) 0/1 0/1 0/2 Middle of Conduit Body (Slides 7 or11) 0/1 0/1 0/2 Caudal Conduit Body/Atrium (Slides 8, 9, 12 or 13) 0/10/1 0/2 Atrium-Stoma-Skin Junction (Slide 18) 0/1 0/1 0/2 Surface ofChronic-Active Inflammation/Detritus Left Ureter Conduit Junction(Slide14) 0/1 1/1 1/2 Right Ureter Conduit Junction (Slide 16) 1/1 1/11/2 Cranial (Ureter) End of Conduit Body (Slides 6 or 10) 1/1 1/1 2/2Middle of Conduit Body (Slides 7 or 11) 1/1 1/1 2/2 Caudal ConduitBody/Atrium (Slides 8, 9, 12 or 13) 1/1 1/1 2/2 Atrium-Stoma-SkinJunction (Slide 18) 1/1 1/1 1/2 Attenuation, Urothelium 1/1 0/1 2/2Hyperplasia, Urothelium 0/1 0/1 0/2 Vacuolation, Urothelium 0/1 0/1 0/2Hemorrhage, Conduit Wall 1/1 0/1 1/2 Scaffold Material, Conduit Wall 0/10/1 2/2 Increased Collagen, Scaffold Wall 1/1 0/1 0/2 Adhered GI Tract(P = present) 0/1 1/1 0/2 Fistula/neutrophil Tract to Intestines 0/1 1/10/2 Acanthosis, skin, stoma 0/1 1/1 1/2 Brown and Hopps Gram Stain Grampositive bacteria in detritus 1/1 1/1 2/2 Gram negative bacteria indetritus 1/1 1/1 2/2

The conduit that developed after surgical implantation of the testarticle consisted of a central lumen that coursed from ureters (cranialend) through the implant and atrium to the stomal opening in the skin ofthe ventral abdomen. The extent of urinary-like tissue regeneration inthe construct groups (Groups 2 and 3) was influenced by duration ofanimal survival postimplantation. Since animals were sacrificed atvarious time points post-implantation, the observed regenerative processwas in different stages and the extent of urinary-like tissue present ineach group varied based on time post-implantation and presence orabsence of SMC.

Ureter-Conduit Junction, Cranial and Mid-Portions of Conduit. Thetypical composition of tissue near the cranial end of the conduitencompassing the ureterconduit junction (UCJ; Sections 14 and 16 in FIG.44) was urothelium overlying a variably-sized submucosa and layers ofsmooth muscle fibers with interspersed connective tissue (FIGS. 27-28).

FIG. 45 shows subgross photographs of animal 6 of Group 3(adipose-derived SMC) in the upper panel and animal 1 of Group 1 in thelower panel.

FIG. 46 shows a photomicrograph (Massons's trichrome stain) of aneo-urinary conduit near the ureter-conduit junction from Group 2 animal4 (blood-derived SMC). The urothelium is evident over a thin submucosaand smooth muscle layers.

FIG. 47 shows a photomicrograph (Massons's trichrome stain) of aneo-urinary conduit near the ureter-conduit junction from Group 3 animal6 (adipose-derived SMC). The urothelium is evident over a thin submucosaand smooth muscle layers.

FIG. 48 shows photomicrographs (Massons's trichrome stain) of amid-conduit wall of animal 1 (Group 1) (left panel) and animal 3 (Group2) (right panel). The mid-conduit wall closest to the lumen was oftenlined by chronic-active inflammation. The wall of the scaffold-only(Group 1) animal is made primarily of blue-stained collagen(reparative), while the wall of the construct on the right is primarilymade up of (regenerative) red-staining spindle cells (presumablymyocytes, fibroblasts, and myofibroblasts).

In the animal that survived to scheduled sacrifice, portions of thecranial and mid conduit were similar in appearance. Where urothelium andsmooth muscle layering were present within the conduit, they weremorphologically similar to the ureters, although the conduit wallthickness was typically greater than that of the ureter, particularlywithin diverticula. Diverticula appeared as bi-compartmental portions ofconduit projecting caudally from the left and right ureteral-conduitjunctions. The typical appearance of urothelium was mildly vacuolatedand variable in thickness. Urothelium thickness varied betweenminimally-to-moderately attenuated (especially within a largediverticulum) and mildly hyperplastic.

Urothelium was present at the ureter-conduit junction (UCJ; Sections 14and/or 16 in FIG. 44) in 5/7 animals (Table 4.19): 1 animal in Group 1(animal 1), 1 animal in Group 2 (animal 4) and 3 animals in Group 3(animals 5, 6, and 7). Urothelium was present at the cranial and midportions (Sections 6 and/or 10 in FIG. 44; Sections 7 and/or 11 in FIG.44) in 1/7 animals: 1 animal in Group 2 (animal 4). Smooth muscle waspresent at the ureter-conduit junction (UCJ; Sections 14 and/or 16 inFIG. 44) in 4/7 animals: 1 animal in Group 1 (animal 1), 1 animal inGroup 2 (animal 4) and 2 animals in Group 3 (animals 5 and 6). Smoothmuscle was present at the cranial and mid portions (Sections 6 and/or 10in FIG. 44 Sections 7 and/or 11 in FIG. 44) in 1/7 animals: 1 animal inGroup 2 (animal 4).

The extent of urinary-like tissue regeneration was dependant upon timepost-implantation. The one animal that survived to scheduled sacrifice(Group 2 animal 4, day 83) had urothelium and smooth muscle present atthe UCJ, cranial and mid portions of the conduit.

The following table 4.19 shows the incidence of Urothelium and SmoothMuscle in UCJ, Cranial and Mid Conduit (Group 1=Scaffold-only, Group2=Blood-derived SMC construct, and Group 3=Adipose-derived SMCconstruct).

TABLE 4.19 Group 1 Group 2 Group 3 Total Finding N = 1 N = 3 N = 3 N = 7Presence of Urothelium UCJ (slides 14 and/or 16) 1/1 1/3 3/3 5/7 Cranial(Ureter) End of Conduit Body 0/1 1/3 0/3 1/7 (Slides 6 and/or 10) Middle(Mid) of Conduit Body 0/1 1/3 0/3 1/7 (Slides 7 and/or 11) Presence ofSmooth Muscle UCJ (slides 14 and/or 16) 1/1 1/3 2/3 4/7 Cranial (Ureter)End of Conduit Body 0/1 1/3 0/3 1/7 (Slides 6 and/or 10) Middle (Mid) ofConduit Body 0/1 1/3 0/3 1/7 (Slides 7 and/or 11)

Caudal Portion of Conduit. The point of transition from conduit toatrium varied between animals because the caudal end of the implantfloated freely within the peritoneal wrapping making the transition fromconduit to atrium difficult to define at necropsy. The typicalcomposition of Sections 8 and 12 (presumed caudal conduit, FIG. 44) wasorganized collagen with associated fibroblasts and/or myofibroblasts.Internal to the collagenous wall and closest to the lumen was a layer ofchronicactive inflammation comprised of loosely arranged collagen,capillaries and abundant neutrophils with fewer lymphocytes andmacrophages. Internal to the inflammation, the lumen was often filledwith detritus comprised of degenerate or necrotic inflammatory cells(primarily neutrophils) and cellular debris with admixed bacterialcolonies. By Brown and Hopps gram stain, bacterial colonies were bothgram positive and gram negative. Most of the gram positive bacteria werecocci; however, both cocci and rod-shaped gram negative and grampositive bacteria were observed. The scaffold-only animal (Group 1) hadonly minimal regeneration near the ureter on one side, and the remainderof the conduit body was almost solely comprised of collagen fibers withminimal fibroblasts. In the animals receiving constructs (Groups 2 and3), regeneration tended to be more extensive, and in other parts of theconduit, the wall was comprised of a mix of collagen, fibroblasts andother spindle cells (presumed myofibroblasts and myocytes).

Atrium and Stoma Portions of Conduit. At the region of the atrium-stomalend of the conduit (Sections 9, 13 and 18 in FIG. 44), the stoma-atriumjunction was visible where the organized collagen and adnexa of thestomal dermis apposed the atrium wall. These sections consisted mainlyof squamous epithelium and chronic-active inflammation/detritus. In 3/7animals (animals 3 and 4 of Group 2, and animal 7 of Group 3), thesquamous epithelium (epidermis) of the skin extended cranially for ashort distance over the atrium. The external surface of the atrium wascomprised of loose connective tissue of peritoneum origin. This externalcovering was the equivalent of a serosal layer and contained nerves,blood vessels, adipose tissue, and some areas of fibrous connectivetissue (collagen fibers and fibroblasts). There was no evidence ofurinary-like tissue regeneration at the atrium-stomal end of the conduit(Sections 9, 13 and 18 in FIG. 44) in any animal.

Other Findings. In 2/7 animals, scaffold material was observed in theconduit wall. This was observed in all levels of the conduit body inGroup 3 animal 5 (day 28), and only in the atriumconduit junction ofGroup 3 animal 6 (day 39).

DISCUSSION. Animals survived 28-83 days. One of 7 animals survived untilscheduled sacrifice (animal 4 of Group 2, 83 days). Six of 7 animalswere sacrificed unscheduled: animal 5 (Group 3) was electivelyeuthanized 28 days post-implantation for histopathological analysis and5 animals were euthanized for poor clinical condition between 38 and 63days post-implantation (Group 1 animal 1; Group 2 animals 2 and 3; andGroup 3 animals 6 and 7).

PCV-2 infection was confirmed in 3/7 animals. Two PCV-2 infected animalswere unscheduled necropsy animals. These included one animal in Group 2(animal 2) euthanized on day 38 and another animal in Group 3 (animal 7)euthanized on day 63. The third animal identified with PCV-2 infection,Group 2 animal 4, survived to scheduled sacrifice (83 days). In these 3animals, there were pathological signs compatible with concurrentinfection with PCV-2 during the course of this study. A macroscopicfinding related to viral infection was skin discoloration in one animal(Group 2 animal 4). Microscopic findings included renal changesconsisting of tubular necrosis/fluid/casts/glomerulonephritis,eosinophilic intracellular inclusions in tubular epithelial cells in thekidney, and vasculitis/perivascular inflammation in the kidneys andureters. These findings are among those commonly reported for pigs withPCV-2 infection. The infection with PCV-2 contributed to the poorclinical health of the animals (e.g. lethargy, diarrhea, loss ofappetite) and subsequent humane early euthanasia. Obstruction of urineflow through the conduit and stoma contributed to the morbidity in 4/6unscheduled death animals. These included Group 1 animal 1, euthanizedon day 47; Group 2 animals 2 and 3, euthanized on days 38 and 40; andGroup 3 animal 6, euthanized on day 39. Obstruction was caused by acombination of the surgical placement of the test article along theabdominal floor of the pig where the weight of abdominal visceracompressed the implant, and the use of peritoneum to form an atriumsegment connecting the conduit to the skin resulting in detritus buildupfrom the external environment and mucus in the urine (normal for swine).Relevant postoperative complications that are inherent in this quadrupedanimal model include: (i) abdominal adhesions leading to potentialfistula formation and (ii) location of the test article placement inrelation to the abdominal organs. When intestines were the adheredorgan, the tunica muscularis of the adhered segment of intestine wasoften diminished or eroded at the point of adhesion, as was the atriumwall. Test articles were placed on the ventral abdomen wrapped withperitoneum and formed a urinary conduit from the ureters to the skinsurface. The test articles were anchored at the cranial end to theureters, but floated freely within the peritoneal wrapping at the caudalend where the peritoneum formed the passage through the abdominal walland onto the skin surface. Smooth muscle and/or urothelium formationoccurred near to the conduit-ureter junctions in 5/7 animals and at thecranial end of the conduit (where the implanted test article was presentwithin the peritoneal wrap) in the one animal surviving to scheduledsacrifice (Group 2 animal 4, day 83). The middle and caudal portions ofthe conduit were comprised of fibrous connective tissue walls withouturothelial covering (except in Group 2 animal 4, in which urothelium andsmooth muscle were observed in the middle level of the conduit).Regeneration of urinary-like tissue was evident as early as day 28, withpresence of urothelium, lamina propria and smooth muscle bundles at theureter-conduit junction (UCJ) in electively euthanized animal 5 (Group3, adipose-derived SMC). The regenerative process at the ureteral end ofthe implant resulted in urinary-like tissue formation that wascomparable among animals receiving a construct implant (Groups 2 and 3).The extent of urinary-like tissue regeneration in the construct groups(Groups 2 and 3) was influenced by duration of animal survivalpost-implantation. The one animal surviving to scheduled sacrifice(Group 2 animal 4, day 83) had urothelium and smooth muscle present inthe UCJ, cranial and mid portions of the conduit in spite of a detectedviral infection. However, the peritoneum-only atrium was insufficient tosupport urinary-like tissue regeneration and the tissue formed in theatrium had a wall comprised of fibrous connective tissue withouturothelial mucosal lining The point of transition from conduit to atriumvaried between animals because the caudal end of the implant floatedfreely within the peritoneal wrapping making the transition from conduitto atrium difficult to define at necropsy. The typical composition ofSections 8 and 12 (presumed caudal conduit, FIG. 44) was organizedcollagen with associated fibroblasts and/or myofibroblasts. Peritoneumconduit without cellular construct appears to be insufficient forurinary-like tissue regeneration, but does serve as a source ofvascularization to NUC implants.

Conclusions

Regeneration of urinary-like tissue was evident as early as 28 days withpresence of urothelium, lamina propria and smooth muscle bundles at theureter-conduit junction (Animal 5, Adipose-derived SMC).

Despite complications from a viral infection and animal model, construct(Scaffold seeded with SMC derived from blood or adipose) implantation(Groups 2 and 3, respectively) resulted in the formation of a conduithaving a urinary-like tissue wall composed of mucosa and smooth musclelayers.

The extent of urinary-like tissue regeneration in the construct groups(Groups 2 and 3) was influenced by duration of animal survivalpost-implantation. The one animal that survived to scheduled sacrificeand was infected with PCV-2 (Group 2 animal 4, day 83) had urotheliumand smooth muscle present at the UCJ, cranial and mid portions of theconduit.

There were no apparent differences observed in the regenerative processwhen scaffolds were seeded with SMC derived from blood or adipose,(Groups 2 and 3, respectively) suggesting equivalence between SMCsources in promoting regeneration.

Animal model anatomy led to the observed complications (i.e., locationof test article in abdomen, peritoneum to form the atrium, compressionof the test article by abdominal contents with formation of adhesionsand fistulas, and detritus buildup that lead to subsequent obstruction).

PCV-2 infection was confirmed in 3/7 animals. Two PCV-2 infected animalswere unscheduled necropsy animals. Obstruction of the urinary flow as aresult of surgical implantation site and abdominal content compressingthe lumen contributed to 4/6 unscheduled deaths. One animal (Group 3animal 5) was electively euthanized at day 28 for histopathologicalevaluation.

Example 5 Evaluation of an Implanted Neo-Urinary Conduit Constructs in aSwine Model

The objective of this study was to evaluate the safety and functionalityof the neo-urinary conduit seeded with autologous smooth muscle cells(SMC) derived from the urinary bladder, adipose, or blood. Additionally,a scaffold-only treatment (not seeded with SMC) was evaluated.

Methods: Neo-Urinary Conduit (NUC) test articles were comprised of ascaffold formed from nonwoven polygycolic acid (PGA) felts andpoly(lactic-co-glycolic acid) polymers (PLGA) with or without autologoussmooth muscle cells (SMC). Unless otherwise indicated, the protocolsfollowed herein are essentially the same as those followed in Example 3.

The study consisted of 32 Gottingen minipigs (16 Females, 16 Males),divided into four groups of 8 animals (4 males and 4 females each group)that were assigned to receive one of the four test articles. Biopsies ofurinary bladder, adipose tissue, and a sample of venous blood wereobtained from animals in Groups 1, 2, and 3 (construct) 6-10 weeks priorto test article implantation. Animals in Groups 1, 2, and 3 wereimplanted with a test article seeded with SMC derived from bladder,adipose, or blood, respectively. Animals in Group 4 were not biopsiedand were implanted with a scaffold-only test article. Animals in Groups1-3 experienced two surgical procedures (biopsy plus test articleimplantation) and Group 4 animals experienced one surgical procedure(test article implantation). Test articles were surgically implantedinto all animals (Groups 1-4) on Day 0 by removing the urinary bladder(radical cystectomy) and diverting the ureters to the inflow end of thetest article. Test articles were placed on the ventral floor of theabdominal cavity and intra-abdominally, having no direct exposure toambient air from the skin stoma which emptied off-midline to the rightcranial abdominal quadrant near the xiphoid. Peritoneum was wrappedaround the test article to provide a vascular source, water-tightness,and to channel urine to an outflow skin stoma. The peritoneal wrapextended approximately 5 to 7 cm beyond the caudal end of the testarticle (located in the intra-abdominal cavity) through the abdominalwall to the skin surface to form a channel for urine outflow; astructure referred to as an “atrium” in this report. Blood and urinesamples were collected at designated time points, analyzed, and resultsrecorded. Imaging (fluoroscopy, ultrasonography, and/or endoscopy) ofthe implants, ureters, and kidneys were performed at designated timepoints during the study. Imaging was also performed as needed inresponse to observed clinical symptoms.

The abdominal cavity was opened and the conduit (the outcome ofimplanting a construct or scaffold-only test article), was visualizedand digitally photographed in situ at the animal facility. The conduitwas removed en bloc with the skin stoma, kidneys, and ureters. Theureters were measured, and then detached from the conduit by transversesectioning 3-4 cm away from the anastomoses. Representative tissuesamples of the entire urinary tract from kidneys to skin stoma, regionallymph nodes, and any other lesions observed grossly were obtained. Alltissue samples were placed in 10% Neutral Buffered Formalin (NBF) for24-48 hours for histological processing and evaluation. Post fixation,tissues were processed routinely to microslides and stained withhematoxylin and eosin (H&E) and Masson's trichrome. Slides wereevaluated microscopically.

Results

Mortality: During definitive surgery, the ureters of animal 30 (Group 4)were perforated, thus preventing test article implantation. The animalwas euthanized on Day 0 and not replaced, reducing the total N ofanimals implanted with test articles (construct or scaffold only) to 31and the N of Group 4 to 7 animals. All other animals were successfullyimplanted with test devices and recovered from surgery. Animals survived6-84 days post-implantation. Twenty-four animals were unscheduled deathsand seven animals survived until the scheduled sacrifice.

Findings related to the Safety of the Construct and Scaffold-onlytreatment groups: Postoperative clinical observations were similaracross groups. The most common of these were stoma flow interruption(31/31 animals), loss of appetite (30/31 animals), soft feces (23/31animals) and body weight loss (21/31 animals). During in-life clinicalobservations, skin lesions consistent with PCV-2 infection wereobserved. Twelve of the 24 unscheduled death animals developed one ormore lesions, consistent with distinct pathological features ascribed toporcine dermatitis and nephropathy syndrome (PDNS), associated withPCV-2 infection. The PCV-2 animals will not be discussed in the resultssection as the PCV-2 condition was deemed an assignable cause ofmorbidity unrelated to the device. Available data for all animals wasobtained (data not shown). Animals were classified as PCV-2-infected ifany of the following were observed: 1) Clinically-observed purple skindiscoloration; 2) Microscopic vasculitis or vasculitis/perivasculitisaffecting the kidney, skin or lung; 3) Kidney findings of tubularnecrosis/fluid/casts/glomerulonephritis, or viral inclusions of tubularepithelial cells; 4) Lymphocyte depletion in lumbar lymph nodes in thepresence of 1, 2, or 3.

By these criteria, porcine PCV-2 infection was confirmed in 12 of the 24unscheduled necropsy animals: 5/8 animals in Group 1, 4/8 animals inGroup 2, 2/8 animals in Group 3, and 1/7 animals in Group 4. Animportant clinical observation in all animals was intermittentobstruction of the outflow of urine with or without atrial, stomal, orstent debris accumulation. Obstruction appeared to have been facilitatedby the placement of the test article in the ventral portion of theabdominal cavity and an anatomic relationship where the weight of theoverlying abdominal organs could lead to conduit closure, adhesion andfistula formation, and renal complications (e.g., dilation,inflammation, and/or infection of ureters or kidney).

Surgical placement of the test article was the same in all studyanimals; therefore, obstruction-related complications had a similarpathobiological mechanism (i.e., abdominal viscera resting on theconduit because of the quadruped anatomy or detritus build up in stoma)in all groups. Obstructions led to significant safety findingsconsisting of hydroureter, hydronephrosis, pyelonephritis, adhesions,and fistula formation. Hydroureter and hydronephrosis were linked tointermittent complete obstruction of the urine outflow. Pyelonephritiswas considered secondary to the debris and detritus build-up andbacterial contamination of the stoma from feces and skin and was mostprevalent in the scaffold-only group (Group 4). The surgical positioningof the test article and multiple surgeries for animals in Groups 1-3(biopsy and test article implantation) promoted the formation ofabdominal and pelvic adhesions. The debridement protocol (use of forcepsin tranquilized animals), urinary flow obstruction, viral infection, andadhesions of intestinal tract to the conduit contributed toenteric-conduit fistula formation.

Findings related to Regeneration: The conduit that developed aftersurgical implantation of the test article consisted of a central lumenthat coursed from ureters (cranial end) through the implant and atriumto the stomal opening in the skin of the ventral abdomen. Thehistological appearance of the conduit wall varied depending uponlocation of sample within the conduit and animal survival time. Urinarytissue-like regeneration characterized by mucosa, submucosa and smoothmuscle with a fibrovascular stroma was observed after construct (Groups1-3) test article implantation regardless of SMC source (i.e., bladder,adipose, or blood). Areas of urinary tract tissue, comprised ofcontinuous urothelium with underlying smooth muscle, were observed in amajority of animals implanted with construct test articles. In contrast,a reparative process was observed following implantation of thescaffold-only test article characterized by an abnormal mucosa supportedby fibrovascular stroma with limited smooth muscle. The extent ofurinary-like tissue regeneration in the construct groups was influencedby duration of animal survival post-implantation.

Conclusions: Seven of 31 (23%) animals completed this study. PCV-2 viralinfection and partial to full urinary obstruction of the urinary flow asa result of surgical implantation site and abdominal content compressingthe lumen contributed to 23/24 unscheduled deaths. An in-life surgicalprocedure-related complication contributed to 1/24 unscheduled death.Safety findings associated with intermittent obstruction of the conduitwere hydronephrosis, hydroureter, pyelonephritis, adhesions, andfistulas.

Healing and regeneration was observed in portions of conduits derivedfrom any of the construct test articles while healing and repair wasobserved in the conduits derived from the scaffold-only test article,demonstrating that construct implantation resulted in the formation of aconduit having a native urinary-like tissue wall composed of mucosa andsmooth muscle layers.

The difference in healing between the construct test articles(regeneration) and scaffold-only test article (repair) contributed to ahigher incidence of significant renal findings observed with thescaffold-only test article, leading to a determination that thescaffold-only test article was unsuitable for further development.

There were no differences observed in the regenerative process andoutcome between construct test articles, suggesting equivalence betweenSMC sources in promoting regeneration.

The objective of this study was to determine the safety andfunctionality of the Neo-Urinary Conduit (NUC) seeded with autologoussmooth muscle cells (SMC) derived from the urinary bladder, adipose orblood. Additionally, a scaffold only treatment (not seeded with SMC) wasevaluated. The goal was to regenerate a tube-like conduit structurecomposed of urinary-like tissue mucosa and wall after surgical removalof the bladder (total cystectomy) and ureteral reimplantation to theinflow end of the test article. The in-life phase of the study lastedabout 5 months.

Experimental Design

Overview. Thirty-two Gottingen minipigs were divided into four groups(4/sex/group). Animals in Groups 1-3 underwent a surgical biopsyprocedure 6-10 weeks prior to test article (construct) implantation toisolate, characterize, and expand the SMC needed to produce a construct.Construct or scaffold only test articles (Groups 1-4) were surgicallyimplanted on Day 0. After surgical removal of the bladder (radicalcystectomy) the ureters were diverted to the inflow end of the testarticle. Test articles were placed on the ventral floor of the abdominalcavity and kept intra-abdominally with no direct exposure to ambient airfrom the skin stoma which emptied off-midline to the right upperabdominal quadrant near the xiphoid. Peritoneum was wrapped around thetest article to provide a vascular source, water-tightness, and tochannel urine to an outflow skin stoma. The peritoneal wrap was alsoextended to form a channel for urine outflow from the caudal end of thetest article (located in the intra-abdominal cavity) through theabdominal wall to the skin surface, a structure referred to as an“atrium” in this report. The test articles were tube-shaped scaffoldsformed from nonwoven polygycolic acid (PGA) felts andpoly(lactic-co-glycolic acid) polymers (PLGA) seeded with autologous SMC(construct) or without SMC (scaffold only). Group 1 animals wereimplanted with constructs seeded with bladder-derived autologous SMC,Group 2 with constructs seeded with adipose-derived autologous SMC,Group 3 with constructs seeded with blood-derived autologous SMC, andGroup 4 animals with scaffolds only. Table 5.1 provides a summary of thestudy design.

TABLE 5.1 No. of Surgical Intended Group Animals Biopsy ProcedureProcedure Postoperative Necropsy No. Treatment M F (Day ~70 to ~40) (Day0) procedures Time Point 1 Autologous 4 4 Removal of CystectomyFluoroscopic and 84 ± 5 days Bladder urinary bladder followed by neo-ultrasonic SMC and adipose urinary conduit examination, 2 Autologous 4 4biopsy, blood implantation with general health adipose collectiontransposition of assessment and SMC ureters to cranial clinicaltreatment 3 Autologous 4 4 end. Whole test as necessary, Blood SMCarticle wrapped clinical pathology 4 Scaffold 4 4 in peritoneum, andurinalysis only (no with a peritoneal cells) transition from end of testarticle through skin to create incontinent stoma.

Table 5.2 provides an overview of the study (TA=Test Article NA=Notapplicable M=Male F=Female N=No Y=Yes; a—Animal found dead, b—Animaldied under anesthesia, c—Ureter perforated at surgery; animal notreplaced, d-Spare animals not utilized in study).

TABLE 5.2 Implant Survival days Cell Adipose Date (Day following SeedingTissue 0)/ Stent implantation/ Cell Density collected Days after RemovalNecropsy Unscheduled Sex Group Source (×10₆) (g) biopsy (Day) Date death(Y/N)  1 F 1 Bladder 40.7 33.0 67 11 81 N  2 F 35.7 35.0 66 12 68 Y  3 F29 33.0 64 11 69 Y  4 F 29 32.0 63 14 28 Y  5 M 24.9 5.0 63 14 44 Y  6 M29 7.0 48 13 83 N  7 M 29 18.0 49 12 42 Y  8 M 29 13.0 51 NA 9 Y  9 F 2Adipose 33.1 29.0 60 5 33 Y 10 F 29 26.0 68 8 30 Y 11 F 29.0 30.0 58 758 Y 12 F 40 35.0 73 5 70 Y 13^(a) M 29 7.0 57 7 82 Y 14 M 29 9.0 55 863 Y 15 M 29 11.0 57 6 81 N 16^(a) M 24 13.0 56 6 63 Y 17 F 3 Blood 2935.0 59 6 48 Y 18 F 24.25 25.0 57 5 80 N 19 F 29 30.0 57 7 70 Y 20 F40.7 40.0 58 5 6 Y 21 M 29 10.0 41 8 73 Y 22^(b) M 29 0.0 56 7 51 Y 23 M29 0.0 57 6 81 N 24 M 29 11.0 42 8 77 Y 25 F 4 Scaffold NA NA 0 9 84 N26 F only; no NA NA 0 9 84 N 27 F autologous NA NA 0 9 20 Y 28^(a) Fcells NA NA 0 7 17 Y 29 M NA NA 0 7 48 Y 30^(c) M NA NA NA NA NA Y 31 MNA NA 0 6 55 Y 32 M NA NA 0 6 31 Y 33^(d) F spares NA NA 26.0 NA NA NASpare 34^(d) M NA 0.0 NA NA NA Spare

Blood and urine samples were collected at designated time points,analyzed and recorded.

Imaging (fluoroscopy, ultrasonography, and/or endoscopy) of theimplants, ureters, and kidneys were performed at designated time pointsduring the study. Imaging was also performed as needed in response toobserved clinical symptoms. Animals survived 6-84 dayspost-implantation. At necropsy, the abdominal cavity was opened, theconduit visualized and photographed before the conduit was removed enbloc with kidneys and ureters. Ureter lengths and widths were measuredand then detached from the conduit by transverse sectioning 3-4 cm awayfrom the anastomoses. Representative sections of the kidneys, ureters,lymph nodes and any other lesions observed grossly were obtained. Alltissue samples were placed in 10% Neutral Buffered Formalin (NBF) for24-48 hours for histological processing and evaluation. Post-fixation,tissues were processed routinely to microslides and stained withhematoxylin and eosin (H&E) and Masson's trichrome. Slides wereevaluated by a board-certified pathologist.

The pathology report appears in the Example below.

Materials and Methods

Test Devices

The test articles were i) a conduit shaped scaffold comprising asynthetic lactide/glycolide polymer seeded with autologousbladder-derived pig smooth muscle cells (24.9×10⁶ cells, 40.7×10⁶ cells,29×10⁶ cells, 24.25×10⁶ cells, or 35.7×10⁶ cells); ii) a conduit shapedscaffold comprising a synthetic lactide/glycolide polymer seeded withautologous blood-derived pig smooth muscle cells (40.7×10⁶ cells or29×10⁶ cells) iii) a conduit shaped scaffold comprising a syntheticlactide/glycolide polymer seeded with autologous adipose-derived pigsmooth muscle cells (24×10⁶ cells, 40×10⁶ cells, 33.1×10⁶ cells, or29×10⁶ cells), and iii) a conduit shaped scaffold comprising a syntheticlactide/glycolide polymer without any cells seeded.

Thirty-two Gottingen minipigs (16 female and 16 male) were received fromMarshall Farms (North Rose, N.Y.) on May 13, 2008. The animals were 7months old. Three additional minipigs (2 females and 1 male) werereceived from the same vendor. These additional animals, which were 6months old, were acquired as spares. Upon receipt, animals were examinedfor signs of disease or injury. Following initial examination, theanimals from the initial shipment were held in quarantine for 6 daysfrom day of receipt and were released for use in the study following aphysical examination. The spare animals were held in quarantine for 4days but were not used for this study.

Identification. Each animal was identified with a unique number that wasindicated by ear tag and on the pen cards.

Initial group assignment was based on average group weight. Assignmentchanges for animals in Groups 1-3 were made based on ex-vivo cellexpansion from harvested tissues and peripheral blood.

Surgical Procedures.

Biopsy/Tissue Collection: For animals in Groups 1-3 (construct testarticles), biopsies of urinary bladder and adipose tissue, as well asvenous blood, were obtained 6-10 weeks prior to Day 0 (implantationprocedure). Group 4 animals (scaffold-only test article) did not undergothese procedures. For tissue biopsy procedures, a midline incision wasmade in the abdomen beginning immediately caudal to the umbilicus.Adipose biopsies of approximately 20-50 grams of soft, pliablesubcutaneous adipose tissue (without connective tissue) were collectedfrom this midline access point, if possible. Bladder biopsies(approximately 2.5 cm×2.5 cm) were collected from the apical dome of theurinary bladder. Collected tissue samples were individually andaseptically transferred to containers with tissue culture media. Thedefect in the bladder was then closed in at least two layers, usingabsorbable suture material. The abdominal incision was closed in layerswith absorbable suture material of an appropriate size. The skin wasclosed in a subcuticular fashion, again using an appropriate size ofabsorbable suture material.

Approximately six 10-ml aliquots of venous blood per animal werecollected in heparinized vacutainers. The vials of blood were thenpackaged in an ice pack-cooled container (˜8° C.).

Cannulation Procedure. On Day 0 (test article implantation), anindwelling catheter was placed within the jugular vein of each animal tofacilitate blood collection. The area surrounding the right jugular veinwas shaved and prepared as described above. All animals were cannulatedwith a sterile 9.0Fr silicon catheter, which was inserted into the rightexternal jugular vein and secured by suture to prevent movement. Anextra-large DaVINCI port was attached and implanted in a subcutaneouspocket.

Test Device Implantation. A midline abdominal incision was made 5 cmcranial to the umbilicus and extended approximately 15 cm caudally. Theperitoneum was identified and then carefully separated from theabdominal space. Care was taken to ensure the tissue remained intact andvascularized.

The urinary bladder was then exposed and carefully emptied of urine,ensuring no urine entered into the abdominal cavity. The arteries andveins supplying the bladder were identified and ligated. The ureterswere identified, stented and carefully transected from the bladder. Theurethra was over-sewn as it was transected. The bladder was thenremoved. The left ureter was carefully separated from the surroundingretroperitoneal fascia (extending cranially) until there was enoughmobility to reach the right side of test article. The right ureter wasdissected free until it reached the inflow (cranial) end of the testarticle. The ureters were sutured onto the test article with 3-0 Vicrylin a simple continuous pattern. Peritoneum was wrapped around the testarticle to provide a vascular source, water-tightness, and to channelurine to an outflow skin stoma. The peritoneal wrap was also extendedapproximately 5-7 cm from the caudal end of the test article (located inthe intra-abdominal cavity) to form a channel for urine outflow throughthe abdominal wall to the skin surface. The peritoneum was sutured with3-0 Vicryl.

Stoma and Stoma Port Procedures. A stoma was created on the ventralabdominal wall lateral to the mammary glands. The peritoneal atriumwithout construct or scaffold-only, was exteriorized and sutured to theskin. Surgical adhesive was placed along the suture line and where theperitoneum exited the body wall. The suture strands connected to thestents were exteriorized through the stoma for future removal. A stomaport (TRACOE®) was placed. Approximately 1-3 months after implantation,a study specific stoma port was used on the remaining animals. For bothtypes, after the stoma ports were secured the abdominal incision wasclosed with non-absorbable Prolene suture. The skin was closed in aroutine fashion.

Monitoring Procedures. Vital signs (oxygen rate, oxygen [O2] saturation,pulse rate, respiration, and body temperature) were monitored atintervals of approximately 20 minutes throughout the procedure.

Postoperative Procedures.

Postoperative Drug Therapy. Concurrent therapy was permissible tomaintain good animal health with the exception of aminoglycosides,quinolones, and corticosteroids. Each drug's identification, dose, routeand frequency of administration were documented and are maintained inthe study file.

During the survival period, clinical symptoms developed related to theprogression of the study. These were addressed by the Facilityveterinarian through drug therapy. The following drugs were utilized.Buprenorphine was used to control pain in the post operative period.Rimadyl (carprofen, an anti-inflammatory) was used post operatively andthen as needed (i.e., post debridement of the stomal area). Excede(ceftiofur) is a long-acting cephalosporin antibiotic. This was usedprophylactically to prevent infection, especially of the urinary tract.Baytril (enrofloxacin) is a fluoroquinolone antibiotic and was used totreat pneumonia and severe UTIs that did not respond to ceftiofur.Reglan (metoclopramide) was used to treat vomiting or constipation.Yobine was used to reverse the effects of the sedative xylazine.

Stent Removal. At various time points from 5-14 days post-implantationsurgery the animals were anesthetized as described above and theureteral stents were removed.

Postoperative Management.

Stoma port. For 2 weeks following implantation, stoma port was evaluateddaily for patency (urine drainage). If urine was not observed to bedripping at examination, the stoma port was flushed with saline toassess patency.

Incision Site. Incision sites were also evaluated daily for 2 weeksfollowing implantation (or until healed) for dehiscence, abnormaldischarge, odor, irritation or any abnormalities.

Debridement: A debridement procedure was initiated because adetritus/caseous material was accumulating in the conduits andpotentially impeding the free flow of urine. Animals on study hadreached 35-52 days post-implantation. For debridement, animals weresedated as described. If the detritus material was larger than the stomaopening, a small incision was made on the stoma to facilitatedebridement. The detritus was visually identified, grasped with forcepsand gently tugged. Once all visible detritus was removed, the stoma wasflushed with saline solution. Stoma incisions were closed with suture(s)and a fresh Study Specific Stoma port was inserted and secured to theanimal with sutures. Animals were allowed to recover as per protocol.

Jugular Port. The jugular port catheter was flushed with injectablesaline and locked with Heparin (100 U/mL, ˜2-3 mL) weekly until Week 4and at each subsequent use to assure patency.

Imaging

Intravenous Pyelography. During Week 8 and prior to necropsy, pyelogramswere obtained following injection of a radiopaque contrast agentdirectly into the renal artery under fluoroscopic guidance. Animals weresedated for the procedure, and access was gained via either the femoralartery or a peripheral vein.

Loopography (Retrograde Pyelography). Five animals [Group 2 animals 9(day 33) and 10 (day 30); Group 3 animals 17 (day 48) and Group 4animals 27 (day 20) and 29 (day 41)] underwent loopography to confirmthe presence of fistulas. Those animals were prepared for fluoroscopicimaging by washing, rinsing and drying the stoma abdominal area. A 3:1mixture of saline: contrast medium was injected via the stoma into theconduit. Fluoroscopic imaging was then performed.

Ultrasonography. Ultrasound imaging of the conduit and kidneys wasperformed at baseline (kidneys only), during Weeks 2, 6, and 10, andprior to necropsy. Animals were anesthetized for this procedure.

Observations.

Health Assessments. From the time of receipt until euthanasia, theanimals were observed twice daily for abnormalities of appearance andbehavior that might indicate adverse effects on health. At each check(performed approximately 8 hours apart), the technician walked throughstudy room to confirm that all animals had eaten and that there wasevidence of urine and fecal output in each cage pan. Physical signs suchas lethargy, emaciation, abnormal vocalization, missing anatomy, andlaceration, as well as abnormal behavioral signs, were noted, ifpresent. Animals were not removed from their cages during these dailyassessments. Any abnormal signs were documented.

Body Weights. Body weights were recorded prior to biopsy (Groups 1-3),prior to implant surgery (Groups 1-4), and prior to necropsy (Groups1-4).

Clinical Pathology.

Blood. Blood samples for analysis of hematology (CBC), coagulation, andserum chemistry parameters were collected prior to biopsy (Groups 1-3)and prior to implant surgery (Group 4), at Weeks 1, 2, 3, 4 and 8, andprior to necropsy via the indwelling port in the jugular vein (Groups1-4). Blood samples for analysis of blood gases were collected prebiopsy(Groups 1-3) and pre-implant surgery (Group 4) via either the jugular orfemoral vein.

The analysis of hematology, coagulation, serum chemistry, and bloodgases were performed as described in Example 3.

Additional Analysis. Immediately prior to euthanasia, heparinized bloodand serum samples were collected from animals 23, 18, 12, 6, 1, and 15.One 10-mL sample of whole blood was collected in a heparin tube.Additional blood was collected in two 4.0-mL serum separation tubes, andserum was separated as described above.

Urine. Two methods were used to collect urine samples at scheduled timepoints (pre-biopsy [Groups 1-3], pre-implant surgery [Group 4] and priorto necropsy): catheterization or a test tube catch method. The methodused was recorded. The desired sample size was at least 3 ml, which wasto be separated into three aliquots in sterile 5-ml tubes. When samplecollection was difficult, the quantitative samples took precedent.Samples were evaluated for qualitative and quantitative parameters asdescribed in Example 3.

Results.

Biopsy. Bladder tissues and blood samples were obtained according toprotocol from all animals in Groups 1, 2, 3 and spares. Adipose tissuebiopsies from all male animals were less than the protocol-definedminimum of 20 grams. However, sufficient SMC were expanded from thecollected tissue to produce constructs for the males assigned to Group2. No adipose tissue could be obtained from three animals (animals 22and 23, and spare animal 34). Animals 22 and 23 were assigned to Group3. Individual weights of adipose biopsy samples are presented in Table5.2.

Implantation. All animals were implanted without incident with twoexceptions. Animal 30 (Male, Group 4), had anatomically small uretersthat were perforated upon attempting to place the stents at test articleimplantation. This animal was euthanized and not replaced. Animal 29(Male, Group 4), also had a small ureteral perforation at test articleimplantation. This animal was successfully implanted and survived thesurgical procedure.

Stent Removal. Individual stent removal data are presented in Table 5.2.Ureteral stents were removed 5-14 days post implantation.

Mortality. Survival days post-implant and final disposition of each ofthe 31 animals implanted with the test article are shown in Table 5.2(above) and Table 5.5 (below). Animal 30 (Group 4) was not implantedwith a test article because of ureteral perforations. The animal waseuthanized and not replaced, reducing the total N to 31 and the N ofGroup 4 to 7 animals. Twenty-four animals were unscheduled deaths andseven animals survived until the scheduled sacrifice. Disposition(mortality classification) of the 24 unscheduled deaths is summarized bytreatment group in Table 5.3.

TABLE 5.3 Mortality Group 1 Group 2 Group 3 Group 4 Total Group N 8 8 87 31 In-life procedure-related 0/8 0/8 1/8 0/7 1/31 death Found dead 0/82/8 0/8 1/7 3/31 Euthanized 6/8 5/8 5/8 4/7 20/31  Total Unscheduled 6/87/8 6/8 5/7 24/31  Necropsies Scheduled Necropsies 2/8 1/8 2/8 2/7 7/31

PCV-2 Associated Mortalities. During in-life clinical observations, skinlesions consistent with PCV-2 infection were observed. Twelveunscheduled death animals developed one or more lesions, consistent withdistinct pathological features ascribed to porcine dermatitis andnephropathy syndrome (PDNS), associated with PCV-2 infection. Animalswere classified as PCV-2-infected according to the criteria of Example3. By these criteria, porcine PCV-2 infection was identified in 12 ofthe 24 unscheduled necropsy animals. These included 5 animals in Group 1(animals 5, 7, 2, 4, and 3); 4 animals in Group 2 (animals 13, 16, 14,and 10); 2 animals in Group 3 (animals 20 and 19); and 1 animal in Group4 (animal 28). The PCV-2 animals will not be discussed in the resultssection as the PCV-2 condition was deemed an assignable cause ofmorbidity unrelated to the device.

Non-PCV-2 Associated Mortalities. A total of 12 unscheduled deaths werenot associated with a detected PCV-2 virus infection (including the 1in-life procedure-related death). The 12 unscheduled deaths notattributable to PCV-2 infection included 1 animal in Group 1 (animal 8);3 animals in Group 2 (animals 11, 9, and 12), 4 animals in Group 3(animals 24, 21, 22, and 17) and 4 animals in Group 4 (animals 32, 31,29, and 27). An important event in the clinical decline of these animalswas obstruction of the outflow of urine and debris through the stoma.Obstruction appeared to have been facilitated by the placement of thetest article in the ventral portion of the abdominal cavity where theweight of the overlying abdominal organs could lead to conduit closure,adhesion and fistula formation, and renal complications (e.g., dilation,inflammation, and/or infection of ureters or kidney). Surgical placementof the test article was the same in all study animals; therefore,obstruction-related complications had a similar pathobiologicalmechanism (i.e. abdominal viscera resting on the conduit because of thequadruped anatomy) in all groups.

Distribution of Unscheduled Deaths by Underlying Findings. A summary ofunderlying findings in the 24 unscheduled deaths is presented in Table5.4.

TABLE 5.4 Mortality Categories Group 1 Group 2 Group 3 Group 4 TotalEvidence of PCV-2 5 4 2 1 12 infection Non-PCV-2 associated 1 3 4 4 12unscheduled deaths Total unscheduled deaths 6 7 6 5 24

Mortality category for each animal implanted with the test articleappears in Table 5.5 (S=scheduled sacrifice; FD=found dead; E=euthanizedfor poor clinical condition; A=died during post-op proceduralcomplication mid-study; SMC=Smooth Muscle Cells; NA=Not applicable;F=Female; M=Male; P=PCV-2 associated mortality; Non-P=Non-PCV-2associated mortality; SURV=survived to scheduled sacrifice.)

TABLE 5.5 Days on Dis- Mortality Animal Sex Group SMC Source Studyposition Group 5 M 1 Bladder 44 E P 2 F 68 E P 8 M 9 E Non-P 4 F 28 E P1 F 81 S SURV 3 F 69 E P 6 M 83 S SURV 7 M 42 E P 13 M 2 Adipose 82 FD P16 M 63 FD P 10 F 30 E P 9 F 33 E Non-P 11 F 58 E Non-P 14 M 63 E P 12 F70 E Non-P 15 M 81 S SURV 18 F 3 Blood 80 S SURV 24 M 77 E Non-P 21 M 73E Non-P 20 F 6 E P 22 M 51 A Non-P 23 M 81 S SURV 19 F 70 E P 17 F 48 ENon-P 26 F 4 Scaffold-only, 84 S SURV 32 M no SMC 31 E Non-P 27 F 20 ENon-P 31 M 55 E Non-P 25 F 84 S SURV 29 M 48 E Non-P 28 F 17 FD P

Findings Pertinent to Safety. Individual and group data for all 31animals implanted with the test article was obtained (data not shown).Discussion is focused on the 7 animals surviving to scheduled necropsyand the 12 non-PCV-2 associated unscheduled deaths. The data isorganized as follows:

Clinical Heath Observations and Post-Surgical Care. Individual clinicalhealth observations and post-surgical care for all animals was observed(data not shown). To distinguish between expected post-operativeclinical observations due to surgical trauma and recovery, and theclinical observations anticipated to be related to obstruction (i.e.,safety signal), the data below is broken into 2 categories: observations<30 days and observations >31 days.

Clinical Heath Observations and Post-Surgical Care for 7 SurvivingAnimals. During the first 30 days post-implantation, debridement was notperformed on any animal. Anorexia (not eating) was observed and stomaport maintenance was performed in all 7 animals. Soft feces (diarrhea)were observed in 3 of 7 animals: 2/2 in Group 1 (animal 1) and 1/2 inGroup 3 (animal 23). These observations are not uncommon following aninvasive surgical procedure (Table 5.6—Pertinent Clinical HealthObservations and Post-Surgical Care by Group of 7 Surviving Animals (<30Days PI)).

TABLE 5.6 Group 1 Group 2 Group 3 Group 4 Total Clinical Observations N= 2 N = 1 N = 2 N = 2 N = 7 Debridement 0/2 0/1 0/2 0/2 0/7 Anorexia(Not Eating) 2/2 1/2 2/2 2/2 7/7 Stoma port Maintenance 2/2 1/1 2/2 2/27/7 Soft Feces (Diarrhea) 2/2 0/1 1/2 0/2 3/7

After day 30 (i.e. day 31-84±5/necropsy), debridement was performed toremove debris/detritus in 4 of 7 animals: 1/2 in Group 1 (animal 1); 1/1in Group 2 (animal 15); and 2/2 in Group 3 (animal 23). Animalsimplanted with the scaffold only (Group 4) did not undergo thedebridement procedure. Anorexia (not eating) was observed in 4 of 7animals: 1/2 in Group 1 (animal 1); 1/1 in Group 2 (animal 15); 1/2 inGroup 3 (animal 18) and 1/2 in Group 4 (animal 26). Stoma portmaintenance was performed on all 7 animals. Soft feces (diarrhea) wereobserved in 5 of 7 animals: 2/2 in Group 1 (animal 1 and 6); 1/1 inGroup 2 (animal 15) and 2/2 in Group 3 (animals 23 and 18) (see Table5.7—Pertinent Clinical Health Observations and Post-Surgical Care byGroup of 7 Surviving Animals; (>30 Days PI)).

TABLE 5.7 Group 1 Group 2 Group 3 Group 4 Total Clinical Observations N= 2 N = 1 N = 2 N = 2 N = 7 Debridement 1/2 1/1 2/2 0/2 4/7 Anorexia(Not Eating) 1/2 1/1 1/2 1/2 4/7 Stoma port Maintenance 2/2 1/1 2/2 2/27/7 Soft Feces (Diarrhea) 2/2 1/1 2/2 0/2 5/7

Clinical Heath Observations and Post-Surgical Care for 12 non-PCV-2Unscheduled Deaths. During the first 30 days post-implantation,debridement was not performed on any animal. Anorexia (not eating) wasobserved in 11 of 12 animals: 1/1 in Group 1 (animal no. 8), 3/3 inGroup 2 (animal nos. 9, 11 and 12), 4/4 in Group 3 (animal nos. 17, 21,24 and 22) and 3/4 in Group 4 (animal nos. 27, 32 and 29). Stoma portmaintenance was performed in all 12 animals. Soft feces (diarrhea) wereobserved in 5 of 12 animals: 1/1 in Group 1 (animal no. 8), 1/3 in Group2 (animal no. 11) and 3/4 in Group 3 (animal nos. 21, 22 and 24). Theseobservations are not uncommon following an invasive surgical procedure(see Table 5.8).

TABLE 5.8 Group 1 Group 2 Group 3 Group 4 Total Clinical Observations N= 1 N = 3 N = 4 N = 4 N = 12 Debridement 0/1 0/3 0/4 0/4  0/12 Anorexia(Not Eating) 1/1 3/3 4/4 3/4 11/12 Stoma port Maintenance 1/1 3/3 4/44/4 12/12 Soft Feces (Diarrhea) 1/1 1/3 3/4 0/4  5/12

Two animals were euthanized before day 30; Group 1 animal no 8 (day 9)and Group 4 animal no. 27 (day 20). Two animals were euthanized at day31 and 33; Group 4 animal no. 32 and Group 2 animal no. 9, respectively.These animals are included in the >30 day analysis. After day 30 (i.e.day 31-84±5/necropsy), debridement was performed to removedebris/detritus in 5 of 12 animals: 2/3 in Group 2 (animal nos. 12 and11) and 3/4 in Group 3 (animal nos. 21, 22 and 24). Animals implantedwith the scaffold only (Group 4) were euthanized or found dead prior toimplementation of the debridement procedure. Anorexia (not eating) wasobserved in 7 of 12 animals: 1/3 in Group 2 (animal no. 11); 4/4 inGroup 3 (animal nos. 17, 21, 22 and 24) and 2/4 animals in Group 4(animal nos. 29 and 31). Stoma port maintenance was performed on 10/12animals: 3/3 in Group 2 (animal nos. 9, 11 and 12), 4/4 in Group 3(animal nos. 21, 22, 17 and 24) and 3/4 in Group 4 (animal nos. 29, 31and 32). Soft feces (diarrhea) were observed in 8 of 12 animals: 3/3 inGroup 2 (animal nos. 9, 11 and 12), 4/4 in Group 3 (animal nos. 17, 21,22 and 24) and 1/4 in Group 4 (animal no. 29). The adverse healthobservations were believed to be a consequence of obstruction (Table5.9—Pertinent Clinical Health Observations and Post-Surgical Care byGroup of 12 non-PCV-2—Unscheduled Deaths (>30 Days PI)).

TABLE 5.9 Group 1 Group 2 Group 3 Group 4 Total Clinical Observations N= 1 N = 3 N = 4 N = 4 N = 12 Debridement 0/1* 2/3{circumflex over ( )}3/4 0/4♦▴ 5/12 Anorexia (Not Eating) 0/1* 1/3{circumflex over ( )} 4/42/4♦▴ 7/12 Stoma Button 0/1* 3/3{circumflex over ( )} 3/4 3/4♦▴ 9/12Maintenance Soft Feces (Diarrhea) 0/1* 3/3{circumflex over ( )} 4/41/4♦▴ 8/12 *Animal no. 8 euthanized on day 9. ♦Animal no. 27 euthanizedon day 20. ▴Animal no. 32 euthanized on day 31. {circumflex over( )}Animal no. 27 euthanized on day 33.

Body Weights. Individual and group body weights for all animals arepresented in Tables 5.10 and 5.11 below.

TABLE 5.10 Weight (Kg) Group Δ: pre Bx % weight Δ pre Δ: pre SX % weightΔ pre ANIMAL No. Group Sex Pre-Bx Pre-Sx →pre Sx Bx →pre Sx Necropsy →NxSX →Nx 1 1 bladder F 12.8 19.8 7.0 55% 13.4 −6.4 −32% 2 1 bladder F 12.918.7 5.8 45% 16.6 −2.1 −11% 3 1 bladder F 14.4 18.7 4.3 30% 13.1 −5.6−30% 4 1 bladder F 11.9 17.9 6.0 50% 15.8 −2.1 −12% 5 1 bladder M 14.016.0 2.0 14% 13.0 −3.0 −19% 8 1 bladder M 14.2 16.0 1.8 13% 11.5 −4.5−28% 6 1 bladder M 15.2 15.9 0.7 5% 16.2 0.3 2% 7 1 bladder M 17.2 20.33.1 18% 17.0 −3.3 −16% Mean 14.1 17.9 3.8 27% 14.6 −3.3 −19% Std Dev 1.61.8 2.3 2.1 2.1 9 2 adipose F 16.2 19.7 3.5 22% 20.2 0.5 3% 10 2 adiposeF 13.1 17.8 4.7 36% 15.5 −2.3 −13% 11 2 adipose F 16.5 18.5 2.0 12% 13.1−5.4 −29% 12 2 adipose F 11.9 18.6 6.7 56% 19.2 0.6 3% 13 2 adipose M17.7 17.4 −0.3 −2% FD NA NA 14 2 adipose M 14.0 14.3 0.3 2% 16.4 2.1 15%15 2 adipose M 15.4 17.1 1.7 11% 15.7 −1.4 −8% 16 2 adipose M 17.6 17.2−0.4 −2% FD NA NA Mean 15.3 17.6 2.3 15% 16.7 −1.0 −5% Std Dev 2.1 1.62.5 2.6 2.7

TABLE 5.11 Weight (Kg) Δ: pre Bx % weight Δ pre Δ: pre SX % weight ΔANIMAL Group No. Group Sex Pre-Bx Pre-Sx →pre-Sx Bx →pre Sx Necropsy →Nxpre SX →Nx 17 3 blood F 13.3 18.6 5.3 40% 17.4 −1.2 −6% 18 3 blood F13.2 16.8 3.6 27% 20.1 3.3 20% 19 3 blood F 13.0 18.0 5.0 38% 12.3 −5.7−32% 20 3 blood F 14.8 20.1 5.3 36% 19.2 −0.9 −4% 22 3 blood M 13.8 17.03.2 23% 16.0 −1.0 −6% 23 3 blood M 13.2 16.6 3.4 26% 18.7 2.1 13% 21 3blood M 14.8 15.8 1.0 7% 10.9 −4.9 −31% 24 3 blood M 16.2 18.1 1.9 12%15.7 −2.4 −13% Mean 14.0 17.6 3.6 26% 16.3 −1.3 −8% Std Dev  1.1 1.4 1.63.3 3.1 25 4 scaffold F NA 16.5 NA NA 17.3 0.8 5% 26 4 scaffold F NA14.0 NA NA 10.9 −3.1 −22% 27 4 scaffold F NA 15.0 NA NA 12.8 −2.2 −15%28 4 scaffold F NA 18.4 NA NA FD NA NA 29 4 scaffold M NA 16.1 NA NA13.0 −3.1 −19% 30 4 scaffold M NA NA NA NA NA NA NA 31 4 scaffold M NA18.2 NA NA 16.3 −1.9 −10% 32 4 scaffold M NA 16.1 NA NA 12.4 −3.7 −23%Mean NA 16.3 NA NA 13.8 −2.2 −16% Std Dev NA 1.6 NA NA 2.5 1.6

Body Weights for 7 Surviving Animals. The body weight of all 7 animalsfluctuated during the study due to post-surgical complications (e.g.,animal model complications leading to obstruction, abdominal adhesions,fistulas and renal complications) (Table 5.12). Although the number ofanimals in each group is small, all animals receiving constructs (Groups1-3) gained weight from pre-biopsy to pre-surgery. All groups exceptGroup 3 lost weight from the time of implant until necropsy.

TABLE 5.12 Weight (Kg)--Average* Δ: Pre- Δ: Pre- % Δ Pre- Surgery→ % ΔPre- Pre- Pre- Biopsy→ Biopsy→ Pre- Pre- Surgery→ Group Biopsy SurgeryPre-Surgery Pre-Surgery Necropsy Necropsy Pre-Necropsy 1 14.0 17.9 3.930% 14.8 −3.1 −15% (N = 2) 2 15.4 17.1 1.7 11% 15.7 −1.4 −8% (N = 1) 313.2 16.7 3.5 27% 19.4 2.7 16% (N = 2) 4 NA 15.3 NA NA 14.1 −1.2 −9% (N= 2)

Body Weights for 12 non-PCV-2 Unscheduled Deaths. The body weight of all12 animals fluctuated during the study due to post-surgicalcomplications (e.g., animal model complications leading to obstruction,abdominal adhesions, fistulas and renal complications) (Table 5.13). Allanimals receiving constructs (Groups 1-3) gained weight from pre-biopsyto pre-surgery. All groups lost weight from the time of implant untilnecropsy, with Group 1 losing the most (N=1) and Group 2 losing theleast amount of weight. (Table 5.13—Average Body Weight by Group for 12Non-PCV-2 Unscheduled Deaths; NA=not applicable).

TABLE 5.13 Weight (Kg)--Average Δ: Pre- % Δ Pre- Δ: Pre- % Δ Pre-Surgery→ Surgery→ Pre- Pre- Biopsy→ Biopsy→ Pre- Pre- Pre- Group BiopsySurgery Pre-Surgery Pre-Surgery Necropsy Necropsy Necropsy 1 14.2 16.01.8 13% 11.5 −4.5 −28% (N = 1) 2 14.9 18.9 4.1 30% 17.5 −1.4  −8% (N =3) 3 14.5 17.4 2.9 20% 15.0 −2.4 −14.2%   (N = 4) 4 NA 16.4 NA NA 13.6−2.7 −17% (N = 4)

Clinical Pathology. Clinical pathology data for individual animals ispresented below (Hematology (CBC); coagulation; serum chemistry; bloodgas; and urinalysis.

Hematology data can be found in Tables 5.14 (White Blood Count (THSN/UL)Ref. Range 11-22); 5.15 (Red Blood Count (MILL/UL) Ref. Range 5-8); 5.16(Hemoglobin (%) Ref. Range 10-16); 5.17 (Hematocrit (%) Ref. Range32-50); 5.18 (MCV (FL) Ref. Range 50-68); 5.19 (MCH (pico gram) Ref.Range 17-21); and 5.20 (PLATELET (THSN/UL) Ref. Range 325-715). Thisdata is based on the individual values of the animals in each group(data not shown).

TABLE 5.14 Base- Week Week Week Week Week Pre- Group Stat line 1 2 3 4 8Necropsy 1 Mean 8.1 21.6 26.6 16.6 23.8 16.5 18.0 SD 1.3 12.5 14.6 5.59.9 3.5 8.1 2 Mean 6.6 15.6 17.0 20.3 18.1 21.9 25.4 SD 2.0 6.9 3.7 6.33.8 9.9 11.7 3 Mean 9.4 14.3 18.8 17.1 19.7 22.4 22.0 SD 2.6 4.0 5.4 4.44.8 7.7 10.6 4 Mean 11.5 14.5 25.2 30.9 25.3 27.3 34.5

TABLE 5.15 Base- Week Week Week Week Week Pre- Group Stat line 1 2 3 4 8Necropsy 1 Mean 6.75 6.47 6.08 6.77 6.81 5.11 5.66 SD 0.95 0.99 1.541.03 0.69 0.33 1.71 2 Mean 6.36 6.64 6.03 6.48 5.89 6.03 5.68 SD 0.411.19 1.21 0.99 0.56 1.24 2.18 3 Mean 6.59 7.39 7.21 6.37 6.42 5.42 5.12SD 0.71 1.26 1.38 1.25 0.83 0.92 1.28 4 Mean 6.45 6.32 6.41 7.10 6.806.11 5.11

TABLE 5.16 Base- Week Week Week Week Week Pre- Group Stat line 1 2 3 4 8Necropsy 1 Mean 11.1 10.9 10.4 11.7 11.7 8.6 9.4 SD 1.7 1.2 2.7 1.9 0.80.9 2.8 2 Mean 10.0 10.5 9.6 10.3 9.4 9.3 9.1 SD 0.9 2.0 1.5 1.2 1.0 1.23.2 3 Mean 10.4 12.0 11.9 10.4 10.5 8.7 8.0 SD 1.2 2.3 2.7 2.1 1.2 1.22.3 4 Mean 9.6 9.5 9.7 10.6 10.1 8.9 7.3 SD 0.7 0.6 1.1 1.9 2.3 3.6 NA

TABLE 5.17 Base- Week Week Week Week Week Pre- Group Stat line 1 2 3 4 8Necropsy 1 Mean 34.3 34.2 32.7 36.8 37.4 27.2 29.0 SD 5.3 3.9 8.2 5.82.5 2.7 8.5 2 Mean 31.0 32.8 30.3 33.1 30.6 30.0 27.9 SD 2.8 5.8 4.7 3.83.4 4.1 10.0 3 Mean 32.3 38.1 36.9 32.2 33.1 27.6 25.6 SD 3.9 7.0 8.06.5 3.5 3.9 7.7 4 Mean 29.7 29.5 29.6 32.4 30.8 28.2 23.4 SD 2.0 1.7 3.55.5 6.9 11.2 NA NA = not applicable, n is too small for SD

TABLE 5.18 Base- Week Week Week Week Week Pre- Group Stat line 1 2 3 4 8Necropsy 1 Mean 51 53 54 54 55 54 51 SD 4 3 3 4 3 4 4 2 Mean 49 49 51 5252 50 49 SD 2 2 3 3 4 3 3 3 Mean 49 52 51 50 52 51 50 SD 3 3 3 3 3 5 3 4Mean 46 47 46 46 45 46 47 SD 2 2 2 2 1 2 NA

TABLE 5.19 Base- Week Week Week Week Week Pre- Group Stat line 1 2 3 4 8Necropsy 1 Mean 16.4 17.0 17.1 17.3 17.2 16.8 16.6 SD 1.3 1.0 1.0 1.31.1 1.6 1.1 2 Mean 15.6 15.8 16.0 16.0 16.0 15.7 16.1 SD 0.6 0.8 0.8 0.91.0 1.2 0.9 3 Mean 15.8 16.2 16.4 16.2 16.4 16.1 15.6 SD 1.0 1.1 1.2 1.21.0 1.5 1.0 4 Mean 15.0 15.0 15.1 14.9 14.9 14.4 14.2

TABLE 5.20 Base- Week Week Week Week Week Pre- Group Stat line 1 2 3 4 8Necropsy 1 Mean 419 404 510 418 379 559 322 SD 89 176 167 171 126 172397 2 Mean 488 407 531 490 489 605 478 SD 61 143 163 151 208 286 435 3Mean 558 434 357 404 453 563 509 SD 130 104 107 115 198 366 388 4 Mean568 473 667 537 429 450 557 SD 126 95 159 199 103 14 NA NA = notapplicable, n is too small for SD

Table 5.21 shows the N-Values by Time Point.

TABLE 5.21 N-Values by Time Point: Base- Week Pre- Group line 1 Week 2Week 3 Week 4 Week 8 Necropsy 1 8 8 7 7 6 4 7 2 8 8 8 8 8 6 6 3 8 8 7 77 5 6 4 8 7 7 6 5 3 2

Coagulation data is provided in Tables 5.22 (Prothrombin Time (seconds)Ref. Range 10-15.5); 5.23 (Activated Partial Thromboplastin Time (s)Ref. Range 18.4-27.7); and 5.24 (Fibrinogen (MG/DL) Ref. Range 100-500).This data is based on the individual values of the animals in each group(data not shown).

TABLE 5.22 Base- Week Week Week Week Week Pre- Group Stat line 1 2 3 4 8Necropsy 1 Mean 11.5 11.8 11.7 12.0 11.5 11.4 16.3 SD 0.8 1.1 1.6 0.70.8 0.7 5.5 2 Mean 11.3 11.4 11.4 10.9 11.3 11.5 12.6 SD 0.8 0.4 0.5 0.40.3 0.4 1.2 3 Mean 11.4 11.2 11.7 10.9 11.4 11.3 12.2 SD 0.7 1.4 0.4 0.40.6 1.3 1.7 4 Mean 11.1 11.7 11.9 13.0 12.2 12.9 12.3 SD 0.3 0.2 0.7 0.90.6 2.0 0.6

TABLE 5.23 Base- Week Week Week Week Week Pre- Group Stat line 1 2 3 4 8Necropsy 1 Mean 34.63 32.18 36.96 0.00 33.62 29.05 22.14 SD 9.47 9.0615.08 0.00 10.19 8.09 7.70 2 Mean 33.88 32.89 39.53 37.20 29.95 20.5720.18 SD 6.80 9.73 12.53 10.81 7.17 3.80 3.93 3 Mean 31.65 24.74 41.2637.40 34.33 22.74 18.92 SD 6.36 5.47 17.12 14.67 14.41 7.61 5.20 4 Mean40.51 41.66 30.29 36.10 30.52 38.00 29.05 SD 2.80 9.16 9.36 7.93 4.0111.10 NA

TABLE 5.24 Base- Week Week Week Week Pre- Group Stat line Week 1 2 3 4 8Necropsy 1 Mean 520 1561 837 1102 656 797 977 SD 155 2237 468 452 368200 530 2 Mean 473 882 752 562 867 925 1218 SD 179 483 246 261 126 327589 3 Mean 578 1140 591 610 1019 859 1339 SD 129 851 299 275 762 5961276 4 Mean 700 1084 1908 1617 1051 1780 1496 SD 427 698 835 589 462 880NA NA = not applicable, n is too small for SD

Table 5.25 shows the N-Values by Time Point.

TABLE 5.25 Week Week Week Week Week Group Baseline 1 2 3 4 8Pre-Necropsy 1 8 8 7 7 6 4 7 2 8 8 8 8 8 6 6 3 8 8 7 7 7 5 6 4 8 7 7 6 53 2

Serum chemistry data is provided in Tables 5.26 (Sodium (MEQ/L) Ref.Range 135-150); 5.27 (Potassium (MEQ/L) Ref. Range 4.4-6.7); 5.28(Chloride (MEQ/L) Ref. Range 93-106); 5.29 (Total Calcium (MEQ/L) Ref.Range 8.6-10.7); 5.30 (Phosphorous (MG/DL) Ref. Range 9.9-11.9); 5.31(AST SGOT (U/L) Ref. Range 0-32); 5.32 (ALT SGPT (U/L) Ref. Range 0-0);5.33 (ALP (U/L) Ref. Range 0-290); 5.34 (Gamma Glutamyl Transferase(U/L) Ref. Range 10-60); 5.35 (Glucose (MG/DL) Ref. Range 85-150); 5.36(BUN (MG/DL) Ref. Range 10-30); 5.37 (Creatinine (MC/DL) 1-2.7); 5.38(Cholesterol (MG/DL) Ref. Range 36-132); 5.39 (Triglyceride (MG/DL) Ref.Range 50-100); 5.40 (Total Bilirubin (MG/DL) Ref. Range 0.0-1.0); 5.41(Albumin (MG/DL) Ref. Range 1.6-4.0); 5.42 (Total Protein (G/DL) Ref.Range 5.5-8.5); 5.43 (Globulin (G/DL) Ref. Range 1.5-6.9); and 5.44(Albumin/Globulin Ratio (Ratio) Ref. Range 0.2-2.7). This data is basedon the individual values of the animals in each group (data not shown).

TABLE 5.26 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 153 139 141 143 141 138 133 SD 15 3 2 5 4 4 15 2 Mean147 141 136 139 139 137 137 SD 11 5 7 6 2 4 4 3 Mean 152 137 131 142 141141 132 SD 12 6 8 3 4 4 11 4 Mean 140 138 136 131 136 135 140 SD 3 5 5 68 5 NA

TABLE 5.27 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 4.4 4.2 4.6 4.8 4.8 4.3 5.6 SD 0.4 0.3 1.0 0.4 1.2 0.51.1 2 Mean 4.2 4.1 4.0 4.7 4.4 4.7 5.0 SD 0.4 0.5 0.5 0.4 0.4 0.4 0.6 3Mean 4.2 4.3 5.6 4.5 4.7 5.1 5.7 SD 0.2 0.9 2.3 0.4 0.3 0.7 1.7 4 Mean3.9 4.1 5.3 4.3 5.0 4.3 4.6 SD 0.3 0.3 0.7 0.9 2.4 1.4 NA

TABLE 5.28 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 111 98 101 101 98 103 97 SD 14 2 2 2 3 4 23 2 Mean 105100 97 99 101 99 99 SD 12 6 5 4 2 5 11 3 Mean 111 96 94 102 100 103 102SD 12 7 7 5 5 4 11 4 Mean 98 97 94 89 94 87 98 SD 4 6 6 7 10 10 NA

TABLE 5.29 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 10.9 9.9 10.0 10.4 10.3 9.9 9.8 SD 0.7 0.4 0.9 0.5 0.50.3 0.8 2 Mean 10.7 9.6 9.8 10.3 9.9 10.0 9.6 SD 0.6 0.5 0.6 0.5 0.5 0.40.6 3 Mean 10.6 10.0 10.3 9.9 10.2 10.3 9.9 SD 0.7 0.6 0.8 0.4 0.5 0.40.9 4 Mean 10.4 10.3 10.3 10.0 10.2 10.6 10.4 SD 0.3 0.7 0.6 0.6 0.7 0.3NA NA = not applicable, n is too small for SD

TABLE 5.30 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 9.5 6.1 6.2 6.8 6.8 6.5 11.6 SD 1.1 0.6 0.9 0.8 1.2 0.85.9 2 Mean 8.6 5.8 6.8 6.9 6.8 6.2 9.0 SD 0.9 1.5 0.8 0.8 0.7 0.4 5.4 3Mean 8.5 7.2 8.4 6.6 6.6 6.8 8.3 SD 1.4 3.0 3.7 1.7 0.8 0.7 1.8 4 Mean7.7 6.6 7.4 7.4 7.6 6.8 6.6 SD 0.5 0.8 0.7 2.2 2.2 2.1 NA

TABLE 5.31 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 108 41 33 22 21 17 301 SD 202 15 14 10 7 5 577 2 Mean 3045 17 17 20 39 76 SD 13 30 3 4 7 17 106 3 Mean 34 44 34 41 27 31 47 SD15 26 10 25 19 17 26 4 Mean 29 18 27 42 110 23 46 SD 19 7 9 38 112 2 NA

TABLE 5.32 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 61 56 48 30 29 23 29 SD 48 19 19 19 6 10 15 2 Mean 35 5133 29 28 22 26 SD 7 5 12 10 12 7 6 3 Mean 63 46 55 53 36 30 24 SD 69 2218 11 11 6 8 4 Mean 34 29 25 21 41 20 19 SD 4 9 4 5 25 3 NA

TABLE 5.33 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 113 76 59 66 88 63 68 SD 32 30 14 23 34 27 34 2 Mean 12987 75 83 76 66 72 SD 33 31 15 31 27 10 45 3 Mean 96 99 63 65 64 62 65 SD26 47 14 26 19 19 57 4 Mean 102 63 54 65 85 81 91 SD 25 16 9 23 24 14 NANA = not applicable, n is too small for SD

TABLE 5.34 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 64 62 47 58 47 48 50 SD 15 16 8 14 5 4 17 2 Mean 65 4746 48 63 47 51 SD 16 8 9 6 15 8 12 3 Mean 59 65 63 49 47 44 50 SD 12 2518 17 16 12 27 4 Mean 68 44 65 47 55 43 84 SD 16 9 28 9 8 5 NA

TABLE 5.35 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 80 87 78 87 90 83 67 SD 19 16 29 17 24 15 33 2 Mean 8775 86 75 74 76 58 SD 15 17 20 11 19 20 25 3 Mean 89 86 103 71 69 72 240SD 22 14 43 7 7 17 390 4 Mean 77 92 96 69 92 100 76 SD 14 9 19 24 32 8NA

TABLE 5.36 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 15 17 18 13 13 50 160 SD 8 10 11 6 2 30 138 2 Mean 12 2916 19 14 27 140 SD 8 14 16 18 14 15 51 3 Mean 14 42 35 13 17 44 91 SD 740 46 10 12 34 59 4 Mean 7 13 30 49 33 76 44 SD 1 4 16 38 35 78 NA

TABLE 5.37 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 1.2 1.3 1.2 1.2 1.2 1.5 4.3 SD 0.4 0.3 0.3 0.2 0.2 0.54.2 2 Mean 1.1 1.6 1.1 1.1 1.0 1.2 2.0 SD 0.3 0.7 0.5 0.2 0.2 0.3 1.1 3Mean 1.3 3.7 1.8 1.0 1.1 1.4 2.0 SD 0.4 5.6 1.3 0.2 0.4 0.4 1.1 4 Mean1.1 1.1 1.5 1.9 1.8 6.3 3.5 SD 0.1 0.1 0.5 1.2 1.0 5.8 NA

TABLE 5.38 Base- Week Week Week Week Pre- Group Stat line 1 2 3 4 Week 8Necropsy 1 Mean 87 69 69 84 79 77 74 SD 38 14 12 13 16 19 38 2 Mean 7180 83 92 79 74 71 SD 27 24 23 28 36 29 26 3 Mean 73 71 65 75 77 73 66 SD35 10 26 16 17 17 24 4 Mean 72 83 87 84 87 120 130 SD 27 31 51 54 41 37NA

TABLE 5.39 Pre- Group Stat Baseline Week 1 Week 2 Week 3 Week 4 Week 8Necropsy 1 Mean 48 46 47 79 76 37 64 SD 49 11 30 57 62 19 24 2 Mean 4047 132 41 25 45 57 SD 35 21 121 30 12 19 11 3 Mean 57 53 40 42 100 53 41SD 55 12 20 17 55 27 25 4 Mean 21 27 34 48 45 97 45 SD 9 9 15 29 26 8 NA

TABLE 5.40 Pre- Group Stat Baseline Week 1 Week 2 Week 3 Week 4 Week 8Necropsy 1 Mean 0.1 0.1 0.1 0.1 0.2 0.1 0.5 SD 0.0 0.1 0.1 0.1 0.1 0.00.5 2 Mean 0.1 0.2 0.1 0.2 0.1 0.1 0.3 SD 0.1 0.1 0.0 0.1 0.1 0.0 0.2 3Mean 0.1 0.3 0.2 0.1 0.0 0.1 0.2 SD 0.1 0.2 0.1 0.1 0.0 0.0 0.1 4 Mean0.1 0.2 0.1 0.1 0.2 0.4 0.1 SD 0.1 0.1 0.0 0.1 0.1 0.2 NA

TABLE 5.41 Pre- Group Stat Baseline Week 1 Week 2 Week 3 Week 4 Week 8Necropsy 1 Mean 4.3 3.6 3.7 3.4 3.3 3.5 3.0 SD 0.4 0.3 0.6 0.3 0.2 0.30.4 2 Mean 4.1 3.5 3.4 3.5 3.4 3.2 3.1 SD 0.3 0.4 0.4 0.3 0.4 0.2 0.6 3Mean 4.2 3.8 3.8 3.5 3.4 3.4 3.0 SD 0.4 0.4 0.3 0.3 0.5 0.3 0.9 4 Mean4.2 3.7 3.8 3.4 3.4 3.0 3.1 SD 0.3 0.4 0.4 0.3 0.1 0.2 NA

TABLE 5.42 Pre- Group Stat Baseline Week 1 Week 2 Week 3 Week 4 Week 8Necropsy 1 Mean 6.3 6.6 7.1 7.7 7.5 7.7 7.8 SD 0.6 0.7 0.8 0.6 0.7 0.80.9 2 Mean 5.7 6.4 6.6 6.9 6.8 7.1 7.0 SD 0.5 0.6 0.8 0.5 0.5 0.7 0.9 3Mean 6.3 6.9 7.3 6.8 7.3 7.7 7.5 SD 0.7 0.6 1.0 0.7 0.7 0.5 1.4 4 Mean6.1 6.3 7.5 7.4 7.7 8.6 8.1 SD 0.4 0.5 0.8 0.7 0.9 0.4 NA

TABLE 5.43 Pre- Group Stat Baseline Week 1 Week 2 Week 3 Week 4 Week 8Necropsy 1 Mean 2.0 3.1 3.4 4.2 4.2 4.2 4.8 SD 0.3 0.5 0.5 0.7 0.6 0.90.7 2 Mean 1.6 2.9 3.2 3.3 3.4 3.8 3.9 SD 0.3 0.4 0.5 0.4 0.3 0.7 0.6 3Mean 2.1 3.1 3.5 3.3 3.9 4.3 4.5 SD 0.3 0.3 0.9 0.5 0.5 0.3 0.6 4 Mean1.9 2.6 3.7 4.0 4.3 5.6 5.0 SD 0.2 0.3 0.6 0.4 0.9 0.4 NA

TABLE 5.44 Pre- Group Stat Baseline Week 1 Week 2 Week 3 Week 4 Week 8Necropsy 1 Mean 2.1 1.2 1.1 0.8 0.8 0.9 0.6 SD 0.3 0.2 0.2 0.2 0.1 0.20.1 2 Mean 2.7 1.2 1.1 1.1 1.0 0.9 0.8 SD 0.5 0.2 0.1 0.1 0.1 0.2 0.2 3Mean 2.0 1.2 1.1 1.1 0.9 0.8 0.7 SD 0.2 0.2 0.2 0.2 0.2 0.0 0.1 4 Mean2.3 1.4 1.1 0.9 0.8 0.5 0.6 SD 0.2 0.1 0.2 0.1 0.1 0.1 NA NA = notapplicable, n is too small for SD

Table 5.45 shows the N-Values by Time Point.

TABLE 5.45 Week Week Week Week Week Group Baseline 1 2 3 4 8Pre-Necropsy 1 8 8 7 7 6 4 7 2 8 8 8 8 8 6 6 3 8 8 7 7 7 5 6 4 8 7 7 6 53 2

Blood gas data is provided in Tables 5.46 (pH); 5.47 (PCO2 (mmHg)); 5.48(PO2 (mmHg)); 5.49 (BEecf (mmol/L)); 5.50 (HCO3 (mmol/L)); 5.51 (TCO2(mmol/L)); 5.52 (SO2 (Percent)); 5.53 (Sodium (mmol/L)); 5.54 (Potassium(mmol/L)); 5.55 (Ionized Calcium (mmol/L)); 5.56 (Glucose (mg/dL)); 5.57(Hematocrit (Percent)), and 5.58 (Hb) below. This data is based on theindividual values of the animals in each group (data not shown).

TABLE 5.46 Group Stat Baseline 1 Mean 7.327 SD 0.046 2 Mean 7.288 SD0.050 3 Mean 7.329 SD 0.044 4 Mean 7.346 SD 0.048

TABLE 5.47 Group Stat Baseline 1 Mean 68.8 SD 11.0 2 Mean 72.1 SD 11.1 3Mean 65.8 SD 3.9 4 Mean 58.6 SD 6.6

TABLE 5.48 Group Stat Baseline 1 Mean 267 SD 201 2 Mean 220 SD 218 3Mean 222 SD 244 4 Mean 287 SD 200

TABLE 5.49 Group Stat Baseline 1 Mean 10 SD 2 2 Mean 8 SD 3 3 Mean 9 SD4 4 Mean 6 SD 3

TABLE 5.50 Group Stat Baseline 1 Mean 35.7 SD 2.8 2 Mean 34.3 SD 3.2 3Mean 34.7 SD 3.2 4 Mean 32.0 SD 2.1

TABLE 5.51 Group Stat Baseline 1 Mean 38 SD 3 2 Mean 37 SD 3 3 Mean 37SD 3 4 Mean 34 SD 2

TABLE 5.52 Group Stat Baseline 1 Mean 93 SD 11 2 Mean 84 SD 19 3 Mean 85SD 13 4 Mean 98 SD 5

TABLE 5.53 Group Stat Baseline 1 Mean 147 SD 13 2 Mean 142 SD 9 3 Mean146 SD 10 4 Mean 137 SD 2

TABLE 5.54 Group Stat Baseline 1 Mean 4.3 SD 0.5 2 Mean 4.0 SD 0.4 3Mean 4.2 SD 0.3 4 Mean 3.9 SD 0.2

TABLE 5.55 Group Stat Baseline 1 Mean 1.39 SD 0.05 2 Mean 1.43 SD 0.07 3Mean 1.39 SD 0.08 4 Mean 1.39 SD 0.04

TABLE 5.56 Group Stat Baseline 1 Mean 80 SD 18 2 Mean 85 SD 13 3 Mean 88SD 20 4 Mean 78 SD 11

TABLE 5.57 Group Stat Baseline 1 Mean 29 SD 3 2 Mean 28 SD 3 3 Mean 28SD 3 4 Mean 27 SD 2

TABLE 5.58 Group Stat Baseline 1 Mean 9.9 SD 1.1 2 Mean 9.6 SD 0.9 3Mean 9.5 SD 0.9 4 Mean 9.3 SD 0.7

Table 5.59 shows the N-Values by Time Point.

TABLE 5.59 Group Baseline 1 8 2 8 3 8 4 8

The quantitative microscopic urine analysis is provided in Tables 5.60(Urine Glucose (MG/DL) Ref. Range) and 5.61 (Urine Total Protein (MG/DL)Ref. Range 0-100). This data is based on the individual values of theanimals in each group (data not shown).

TABLE 5.60 Group Stat Baseline Pre-Necropsy 1 Mean 12.9 3.3 SD 9.6 3.9 2Mean 9.8 2.5 SD 3.6 2.1 3 Mean 11.9 38.0 SD 7.0 39.2 4 Mean 13.1 2.0 SD25.0 NA

TABLE 5.61 Group Stat Baseline Pre-Necropsy 1 Mean 38.5 278.0 SD 38.3575.2 2 Mean 25.6 24.7 SD 21.0 54.8 3 Mean 34.5 20.5 SD 45.2 33.2 4 Mean9.3 82.5 SD 11.4 NA

Table 5.62 provides the individual Profiles of the Bacteria (/HPF) Ref.Range for each animal.

TABLE 5.62 ANIMAL # Group Sex Baseline Pre-Necropsy 1 1 F TRACE 3+ 2 1 FTRACE 1+ 3 1 F TRACE — 4 1 F TRACE — 5 1 M TRACE 1+ 8 1 M TRACE — 6 1 MTRACE Trace 7 1 M TRACE — 9 2 F TRACE — 10 2 F TRACE — 11 2 F TRACE none12 2 F TRACE — 13 2 M TRACE — 14 2 M TRACE 1+ 15 2 M TRACE Trace 16 2 MTRACE — 17 3 F TRACE — 18 3 F TRACE 2+ 19 3 F TRACE — 20 3 F TRACE — 223 M TRACE — 23 3 M TRACE Trace 21 3 M TRACE none 24 3 M TRACE Trace 25 4F TRACE Trace 26 4 F TRACE Trace 27 4 F TRACE — 28 4 F TRACE — 29 4 MTRACE — 30 4 M TRACE — 31 4 M TRACE — 32 4 M TRACE —

Qualitative urine analysis was also performed for each animal: SpecificGravity (g/mL) Ref. Range 1.001-1.035; Blood (Ery/μL) Ref. Range Neg.(non-Haemolyzed); Trace-Spur (Haemolyzed); pH Ref. Range 4.6-8.0;Protein (mg/dL) Ref. Range <150 mg; Urobilinogen (mg/dL) Ref. Range ≦1;Nitrates (˜Leu/μL) Ref. Range neg; and Leukocytes (˜Leu/μL) Ref. Rangeneg (data not shown).

Urine bacterial culture and sensitivity results were obtained. Bacteriawas cultured from two animals. Animal 18 was found to have ESCHERICHIACOLI—greater than 100,000 organisms per ml; MORGANELLA MORGANIT—greaterthan 100,000 organisms per ml ENTEROCOCCUS SPECIES-2+ ENTEROCOCCUSSPECIES 2+ second strain. Animal 32 was found to have PROTEUSMIRABILIS—greater than 100,000 organisms per ml strain 1 PROTEUSMIRABILIS—greater than 100,000 organisms per ml strain 2 NON-ENTERICGRAM NEG ROD unable to speciate—10,000-50,000 organisms per ml.

Hematology for 7 Surviving Animals. Postoperative blood collection forhematology revealed the development of leukocytosis for all groups.Leukocyte counts for the construct groups (Groups 1-3) rose frombaseline to week 2-4 and then plateaued or decreased thereafter;however, leukocyte counts for the scaffold-only group (Group 4)continued to increase until necropsy (Table 5.63 Average WBC Count byGroup for 7 Surviving Animals (reference range 11-22 THSN/UL)).

TABLE 5.63 Week Week Week Week Pre- Group Baseline Week 1 2 3 4 8Necropsy 1 (N = 2) 7.3 29.4 47.6 23.6 24.0 19.1 13.2 2 (N = 1) 5.7 10.317.9 31.4 15.1 17.7 12.3 3 (N = 2) 9.5 15.6 17.0 12.9 22.0 19.6 13.2 4(N = 2) 10.3 11.9 14.8 28.1 25.4 27.6 34.5

Red blood cell counts (RBCs) were similar across all groups (Table5.64). At necropsy, mean RBC was within the reference range for allgroups except Group 3. (Table 5.64—Average RBC Count by Group for 7Surviving Animals (reference range 5-8 MILL/UL).

TABLE 5.64 Week Week Week Week Pre- Group Baseline Week 1 2 3 4 8Necropsy 1 (N = 2) 6.8 6.6 5.3 5.8 6.9 5.4 6.4 2 (N = 1) 6.3 6.2 5.7 5.85.7 5.6 5.9 3 (N = 2) 6.4 6.7 6.4 6.5 6.8 5.5 4.7 4 (N = 2) 6.2 6.4 6.27.5 7.0 6.8 5.1

Hematocrit values for all groups were below or at the low end ofreference at baseline and fluctuated during the study, with values inthe lower limit of the reference range at week 8 and necropsy (Table5.65—Average Hematocrit by Group for 7 Surviving Animals (referencerange 32-50%)).

TABLE 5.65 Week Week Week Week Pre- Group Baseline Week 1 2 3 4 8Necropsy 1 (N = 2) 34.0 35.0 27.9 30.5 37.1 28.2 30.8 2 (N = 1) 31.032.1 30.5 31.4 31.0 29.4 28.8 3 (N = 2) 32.0 36.0 34.4 33.9 36.2 28.624.1 4 (N = 2) 29.3 30.6 29.0 35.0 32.2 32.1 23.4

Hematology for 12 non-PCV2 Unscheduled Deaths. Postoperative bloodcollection for hematology revealed the development of leukocytosis forall groups (Table 5.66—Average WBC Count by Group for 12 non-PCV2Unscheduled Deaths (reference range 11-22 THNS/UL).

TABLE 5.66 Week Week Week Week Pre- Group Baseline Week 1 2 3 4 8Necropsy 1 (N = 1)* 7.6 42.4 NA NA NA NA NA 2 (N = 3) 5.4 11.9 19.1 21.116.2 31.0 23.8 3 (N = 4) 10.0 13.4 17.5 18.5 18.2 28.5 24.3 4 (N = 4)11.8 15.6 27.6 32.4 25.3 26.8 NA NA = Not applicable *Animal no. 8euthanized on day 9.

Red blood cell counts (RBCs) were within normal ranges and similaracross all groups (Table 5.67—Average RBC Count by Group for 12 non-PCV2Unscheduled Deaths (reference range 5-8 MILL/UL).

TABLE 5.67 Week Week Week Week Pre- Group Baseline Week 1 2 3 4 8Necropsy 1 (N = 1)* 6.6 8.4 NA NA NA NA NA 2 (N = 3) 6.5 7.1 6.9 7.4 6.36.5 6.4 3 (N = 4) 6.3 6.9 7.4 5.7 6.2 5.6 5.6 4 (N = 4) 6.5 6.4 6.5 6.96.7 4.7 NA NA = Not applicable *Animal no. 8 euthanized on day 9.

Hematocrit values for the scaffold only group (Group 4) showed asignificant drop in hematocrit values at week 8 (Table 5.68—AverageHematocrit by Group for 12 non-PCV2 Unscheduled Deaths (reference range32-50%).

TABLE 5.68 Week Week Week Week Pre- Group Baseline Week 1 2 3 4 8Necropsy 1 (N = 1)* 30.1 41.7 NA NA NA NA NA 2 (N = 3) 31.5 34.4 33.836.4 31.0 31.8 30.2 3 (N = 4) 30.8 35.6 37.6 28.9 31.5 28.8 28.1 4 (N =4) 29.6 29.2 29.2 31.1 29.9 20.5 NA NA = Not applicable *Animal no. 8euthanized on day 9.

Coagulation for 7 Surviving Animals. Coagulation panels revealedelevated fibrinogen for all groups throughout the study. The scaffoldonly group (Group 4) had the highest concentration and greatestelevation in fibrinogen from baseline (Table 5.69—Average Fibrinogen byGroup for 7 Surviving Animals (reference range 100-500 mg/dL MG/DL).

TABLE 5.69 Week Week Week Week Pre- Group Baseline Week 1 2 3 4 8Necropsy 1 (N = 2) 673 968 1606 1076 344 785 789 2 (N = 1) 394 1116 381947 1069 794 833 3 (N = 2) 529 1148 502 716 629 1052 1027 4 (N = 2) 419546 1704 1469 868 1297 1496

The activated partial thromboplastin time was elevated or high normalthroughout the study and related to the higher fibrinogen concentration(Table 5.70—Average APTT by Group for 7 Surviving Animals (referencerange 18.4-27.2 Seconds).

TABLE 5.70 Week Week Week Week Pre- Group Baseline Week 1 2 3 4 8Necropsy 1 (N = 2) 44.15 30.85 27.05 38.60 34.00 29.25 16.30 2 (N = 1)29.00 23.30 45.00 29.50 28.30 25.50 16.20 3 (N = 2) 28.25 20.85 36.5028.95 33.00 24.05 24.30 4 (N = 2) 44.35 41.00 33.40 38.00 33.75 44.0529.05

Coagulation for 12 non-PCV-2 Unscheduled Deaths. Coagulation panelsrevealed elevated fibrinogen for all groups throughout the study. Thescaffold only group (Group 4) had the highest concentration and greatestelevation in fibrinogen from baseline (Table 5.71—Average Fibrinogen byGroup for 12 non-PCV-2 Unscheduled Deaths (reference range 100-500MG/DL).

TABLE 5.71 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 (N = 1)* 282 7056 NA NA NA NA NA 2 (N = 3) 406 1007 921567 899 918 1531 3 (N = 4) 603 682 518 603 1253 1051 1859 4 (N = 4) 8691432 1886 1691 1172 2745 NA NA = Not applicable *Animal no. 8 euthanizedon day 9.

The activated partial thromboplastin time generally was elevated or highnormal throughout the study for all groups. (Table 5.72—Average APTT byGroup for 12 non-PCV-2 Unscheduled Deaths (reference range 18.4-27.7Seconds).

TABLE 5.72 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 (N = 1)* 18.90 32.40 NA NA NA NA NA 2 (N = 3) 35.03 31.2031.90 41.15 31.20 16.80 22.77 3 (N = 4) 31.95 27.45 48.13 42.88 33.8827.00 17.07 4 (N = 4) 39.88 42.88 29.60 35.15 28.37 25.90 NA NA = Notapplicable *Animal no. 8 euthanized on day 9.

Serum Chemistry for 7 Surviving Animals.

Increases in BUN are shown in Table 5.73—Average BUN by Group for 7Surviving Animals (reference range 10-30 MG/DL.

TABLE 5.73 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 (N = 2) 12 22 26 17 13 46 89 2 (N = 1) 8 34 9 7 6 44 2143 (N = 2) 12 38 30 21 15 48 42 4 (N = 2) 8 11 23 45 25 34 44

Increases in creatinine are shown in Table 5.74—Average Creatinine byGroup for 7 Surviving Animals (reference range 1-2.7 MC/DL),

TABLE 5.74 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 (N = 2) 1.1 1.4 1.2 1.1 1.2 1.3 1.9 2 (N = 1) 0.9 0.9 0.80.8 0.7 0.9 1.5 3 (N = 2) 1.4 2.3 1.5 1.3 1.3 1.6 1.2 4 (N = 2) 1.1 1.21.2 2.5 1.9 3.1 3.2

Increases in potassium are shown in Table 5.75—Average Potassium byGroup for 7 Surviving Animals (reference range 4.4-6.7 MEQ/L).

TABLE 5.75 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 (N = 2) 4.1 4.4 4.5 5.1 6.3 4.1 6.2 2 (N = 1) 4.6 3.2 4.44.1 4.4 4.4 4.8 3 (N = 2) 4.3 3.3 5.2 4.2 5.1 5.2 5.2 4 (N = 2) 3.7 3.94.9 4.2 3.6 3.8 4.6

Increases in total protein are shown in Table 5.76—Average Total Proteinby Group for 7 Surviving Animals (reference range 5.5-8.5 G/DL).

TABLE 5.76 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 (N = 2) 6.2 6.5 6.8 7.3 7.2 7.1 7.8 2 (N = 1) 5.0 5.8 6.26.7 6.7 6.8 7.2 3 (N = 2) 6.5 7.1 7.4 7.4 7.7 8.0 8.3 4 (N = 2) 6.2 6.67.5 7.5 7.8 8.7 8.1

Decreases in sodium are shown in Table 5.77—Average Sodium by Group for7 Surviving Animals (reference range 135-150 MEQ/L).

TABLE 5.77 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 (N = 2) 154 137 143 140 144 138 141 2 (N = 1) 147 131 145144 140 133 140 3 (N = 2) 146 139 136 139 144 142 140 4 (N = 2) 138 139135 129 138 135 140

Decreases in phosphorous are shown in Table 5.78—Average Phosphorous byGroup for 7 Surviving Animals (reference range 9.9-11.9 MG/DL).

TABLE 5.78 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 (N = 2) 8.9 6.0 5.5 7.0 7.9 6.5 6.4 2 (N = 1) 8.3 3.1 7.96.2 6.9 6.4 5.7 3 (N = 2) 7.6 6.4 7.7 6.9 7.2 7.3 7.1 4 (N = 2) 7.5 6.67.5 7.5 6.8 5.7 6.6

Decreases in albumin are shown in Table 5.79—Average Albumin by Groupfor 7 Surviving Animals (reference range 1.6-4.0 MG/DL).

TABLE 5.79 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 (N = 2) 4.0 3.3 3.2 3.1 3.1 3.4 3.1 2 (N = 1) 3.8 2.9 3.23.3 3.2 3.4 3.9 3 (N = 2) 4.3 4.0 4.0 3.8 3.7 3.5 3.6 4 (N = 2) 4.4 4.04.0 3.5 3.4 3.1 3.1

The increases and decreases shown in Tables 5.73 to 5.79 were observedin all groups. In general, profiles for animals receiving constructimplants (Groups 1-3) appear to be similar in magnitude and temporallyacross; however, the scaffold-only group (Group 4) appears to haveearlier serum changes of greater magnitude that suggest more advancedand serious renal deterioration.

The serum chemistry for 12 non-PCV-2 Unscheduled Deaths was alsoexamined.

Increases in BUN are provided in Table 5.80 Text Table 34: Average BUNby Group for 12 non-PCV-2 Unscheduled Deaths (reference range 10-30MG/DL).

TABLE 5.80 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 6 8 NA NA NA NA NA (N = 1)* 2 12 28 24 19 9 35 124 (N =3) 3 11 22 42 11 21 32 126 (N = 4) 4 6 13 31 51 39 162 NA (N = 4) NA =Not applicable *Animal no. 8 euthanized on day 9.

Increases in creatinine are provided in Table 5.81—Average Creatinine byGroup for 12 non-PCV-2 Unscheduled Deaths (reference range 1-2.7 MC/DL).

TABLE 5.81 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 1.0 0.9 NA NA NA NA NA (N = 1)* 2 1.1 1.7 1.3 1.1 1.0 1.42.0 (N = 3) 3 1.0 1.4 2.0 0.9 1.2 1.3 2.6 (N = 4) 4 1.1 1.1 1.3 1.6 1.712.7 NA (N = 4) NA = Not applicable *Animal no. 8 euthanized on day 9.

Increases in potassium are provided in Table 5.82—Average Potassium byGroup for 12 non-PCV-2 Unscheduled Deaths (reference range 4.4-6.7MEQ/L).

TABLE 5.82 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 4.2 3.9 NA NA NA NA NA (N = 1)* 2 3.8 4.2 3.8 4.8 4.4 5.05.3 (N = 3) 3 4.2 4.5 6.2 4.6 4.5 5.3 5.8 (N = 4) 4 4.1 4.2 5.3 4.4 6.05.4 NA (N = 4) NA = Not applicable; *Animal no. 8 euthanized on day 9.

Increases in phosphorous are provided in Table 5.83—Average Phosphorousby Group for 12 non-PCV-2 Unscheduled Deaths (reference range 9.9-11.9MG/DL).

TABLE 5.83 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 10.2 5.9 NA NA NA NA NA (N = 1)* 2 9.3 5.8 6.9 6.8 6.96.5 11.1 (N = 3) 3 8.6 6.4 9.3 6.5 6.4 6.8 9.4 (N = 4) 4 7.5 6.9 7.1 7.38.2 9.2 NA (N = 4) NA = Not applicable *Animal no. 8 euthanized on day9.

Increases in total protein are provided in Table 5.84—Average TotalProtein by Group for 12 non-PCV-2 Unscheduled Deaths (reference range5.5-8.5 G/DL).

TABLE 5.84 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 (N = 1)* 6.1 6.2 NA NA NA NA NA 2 (N = 3) 6.0 6.8 7.2 7.37.2 7.7 7.6 3 (N = 4) 6.0 6.9 7.2 6.5 7.2 7.6 7.1 4 (N = 4) 6.1 6.3 7.47.3 7.6 8.4 NA NA = Not applicable *Animal no. 8 euthanized on day 9.

Decreases in sodium are provided in Table 5.85—Average Sodium by Groupfor 12 non-PCV-2 Unscheduled Deaths (reference range 135-150 MEQ/L).

TABLE 5.85 Group Baseline Week 1 Week 2 Week 3 Week 4 Week 8Pre-Necropsy 1 140 139 NA NA NA NA NA (N = 1)* 2 (N = 3) 152 145 130 139138 138 135 3 (N = 4) 149 142 130 142 140 142 131 4 (N = 4) 142 137 139132 135 134 NA NA = Not applicable *Animal no. 8 euthanized on day 9.

Decreases in albumin are provided in Table 5.86—Average Albumin by Groupfor 12 non-PCV-2 Unscheduled Deaths (reference range 1.6-4.0 MG/DL).

TABLE 5.86 Week Week Week Week Pre- Group Baseline 1 2 3 4 Week 8Necropsy 1 4.3 3.5 NA NA NA NA NA (N = 1)* 2 4.2 3.7 3.6 3.7 3.6 3.4 3.2(N = 3) 3 4.0 3.8 3.8 3.5 3.4 3.4 2.7 (N = 4) 4 4.2 3.7 3.7 3.4 3.4 2.8NA (N = 4) NA = Not applicable *Animal no. 8 euthanized on day 9.

The increases and decreases in Tables to 5.80 to 5.86 were observed inall groups. In general, profiles for animals receiving constructimplants (Groups 1-3) appear to be similar in magnitude and temporallyacross; however, the scaffold-only group (Group 4) appears to haveearlier serum changes of greater magnitude that suggest more advancedand serious renal deterioration.

Blood Gas Data for 7 Surviving Animals and 12 non-PCV-2 UnscheduledDeaths. All blood gas data were within normal limits for all 19 animals.

Urinalysis for 7 Surviving Animals and 12 non-PCV-2 Unscheduled DeathsFor the 7 animals surviving to scheduled necropsy, the average urineprotein values were within normal ranges of 0-100 mg/dL for all groups,although the scaffold only group (Group 4) had the highest value atnecropsy (Table 5.87). Due to insufficient data available at necropsyfor the 12 non-PCV-2 unscheduled deaths, comparisons could not be made.Table 5.87 provides the Average Urine Protein by Group for 7 SurvivingAnimals.

TABLE 5.87 Group Baseline Pre-Necropsy 1 (N = 2) 22 23 2 (N = 1) 15 12 3(N = 2) 19 47 4 (N = 2) 2 83

Imaging. Intravenous pyelograms (IVPs) and loopograms (retrogradepyelograms) were performed for each animal. The IV Pyelogramvisualization of one or both ureters and kidneys was inconsistent. Insome cases kidneys, ureters and conduit were visible (radiopaque) whilein other cases visibility was very limited or none. Due to the nature ofthe study a dilute contrast solution had to be used which resulted inlimited visualization.

IVP for 7 Surviving Animals. In 6/7 animals (nos. 1, 6, 15, 23, 25 and26), the Week 8 pyelogram was possible. On Week 8, the pyelogram foranimal no. 18 (Group 3) was (inadvertently) not obtained. The impact ofthis deviation, however, is limited to the single missing pyelogram foranimal no. 18 (Group 3). Preeuthanasia pyelograms on all 7/7 animalswere performed.

Representative pyelogram images are provided. FIG. 49 shows pyelogramimages for animal 6 of Group 1 at week 8 (A) and pre-necropsy (B). FIG.50 shows pyelogram images for animal 15 of Group 2 at week 8 (A) andpre-necropsy (B). FIG. 51 shows a pyelogram image for animal 9 of Group2 at week 5 (loopogram: bowel illumination is indicative of a fistula).FIG. 52 shows a pyelogram image for animal 17 of Group 3 at week 7(loopogram: bowel visualization is indicative of a fistula). FIG. 53shows a pyelogram image for animal 21 of Group 3 at week 8 (bothkidneys; respective ureters and conduit illuminated). FIG. 54 shows apyelogram image for animal 24 of Group 2 at week 8 (one kidney andureter visible). FIG. 55 shows pyelogram images for animal 23 of Group 2at week 8 (A) and pre-necropsy—conduit with stoma port (B). FIG. 56shows pyelogram images for animal 25 of Group 4 at week 8 (A) andpre-necropsy (B). FIG. 57 shows pyelogram images for animal 26 of Group4 at week 8 (A) and pre-necropsy (B). FIG. 58 shows pyelogram images foranimal 29 of Group 4 at week 4—ureters visible (A) and a week 4loopogram—bowel fistula (B).

IVP for 12 non-PCV-2 Unscheduled Deaths. These animals did not surviveto the specified pyelogram. Several pre-necropsy pyelograms were notperformed due to ill health, presence of conduit/bowel fistula and earlyeuthanasia.

Loopograms (Retrograde Pyelograms) for 7 Surviving AnimalsRepresentative Loopograms are presented as described above. Of the 7surviving animals one animal was evaluated for enteric-conduit fistulaformation. The loopogram was performed on animal no. 25 during Week 3.The conduit was confirmed to be free of fistulas. Animal was recoveredand continued on study to successful completion.

Loopograms (Retrograde Pyelograms) for 12 non-PCV-2 Unscheduled DeathsEnteric-conduit fistulas were identified during the in-life position ofthe study in 4 of the 12 non-PCV-2 unscheduled death animals: Loopograms(retrograde pyelograms) were performed at varied timepoints to verifythe presence of fistulas in Group 2 animal No. 9 (day 33); Group 3animal No. 17 (day 48) and in Group 4 Animal Nos. 27 (day 20) and 29(day 41). These animals were euthanized and necropsied at 20 to 48 dayspost-implantation.

Ultrasound for 7 Surviving Animals Individual and group ultrasonographydata are presented in Appendix 11. Ultrasonography on the kidneys overthe course of the study showed increases in surface area (length×width)indicating kidney changes (hydronephrosis) in all treatment groups.Group averages were similar over time for all groups for the right andleft kidney (Table 5.88—Average Kidney Surface Area by Group for 7Surviving Animals (L×W, cm2).

TABLE 5.88 Group Baseline Week 2 Week 6 Week 10 Pre-Necropsy Left Kidney1 (N = 2) 26.7 19.8 21.3 41.2 29.2 2 (N = 1) 16.2 18.4 27.8 21.0 39.3 3(N = 2) 14.3 16.1 19.2 26.5 38.3 4 (N = 2) 14.7 15.2 23.5 39.8 17.1Right Kidney 1 (N = 2) 22.1 20.3 26.0 35.8 23.7 2 (N = 1) 15.4 14.2 25.425.4 22.9 3 (N = 2) 13.4 13.5 13.5 43.2 24.8 4 (N = 2) 17.0 18.7 28.527.3 27.9

Ultrasonography on the implant showed a decrease in wall thickness overtime in all of the construct groups (Groups 1-3). The scaffold onlygroup (Group 4) showed a slight increase in thickness (Table5.89—Average Conduit Wall Thickness by Group for 7 Surviving Animals(cm).

TABLE 5.89 Group Week 2 Week 6 Week 10 Pre-Necropsy 1 (N = 2) 0.3270.227 0.152 0.181 2 (N = 1) 0.172 0.169 0.099 0.114 3 (N = 2) 0.3330.239 0.169 0.145 4 (N = 2) 0.196 0.248 0.179 0.222

Ultrasound for 12 non-PCV-2 Unscheduled Deaths. Individual and groupultrasonography data are presented in Table 5.90—Ultrasonography:Summary Results Kidney Measurements (cm)).

TABLE 5.90 Pre-Biopsy Week 2 Week 6 Right Left Right Left Right LeftGroup Stat Len Wid Len Wid Len Wid Len Wid Len Wid Len Wid 1 Mean 6.292.93 6.08 2.92 6.68 3.17 6.88 3.12 7.14 3.38 6.49 3.19 Std. Dev 1.090.40 1.00 0.63 0.69 0.30 1.23 0.25 0.39 0.26 0.89 0.34 2 Mean 5.36 2.735.66 2.70 5.98 2.83 5.96 2.79 6.66 3.77 7.36 3.91 Std. Dev 0.79 0.290.92 0.33 0.93 0.30 0.77 0.40 0.66 0.50 0.56 0.55 3 Mean 6.08 2.91 6.312.99 6.02 2.99 5.87 2.99 6.40 2.97 7.69 3.38 Std. Dev 0.84 0.52 0.950.41 0.85 0.44 0.80 0.14 1.21 0.48 1.58 0.55 4 Mean 5.83 2.62 5.93 2.586.32 3.89 5.37 2.72 7.95 4.05 7.66 3.99 Std. Dev 0.74 0.26 0.42 0.211.78 1.30 0.88 0.37 1.98 1.58 1.51 0.43 Week 10 Pre-Necropsy Right LeftRight Left Group Stat Len Wid Len Wid Len Wid Len Wid 1 Mean 8.78 5.198.11 4.75 7.25 3.27 7.63 3.76 Std. Dev 1.31 0.90 1.02 0.64 0.79 0.190.26 0.29 2 Mean 8.56 4.06 7.94 3.57 7.40 3.53 8.16 3.90 Std. Dev 0.630.78 0.64 0.71 2.50 0.89 1.97 0.52 3 Mean 9.30 5.16 8.01 4.37 8.58 4.517.01 3.30 Std. Dev 1.77 0.47 1.51 1.33 4.32 2.41 2.47 1.22 4 Mean 7.843.35 8.63 4.56 7.01 3.92 5.55 2.91 Std. Dev NA NA NA NA NA NA NA NA Len= length Wid = width NA = not applicable, n is too small for SD

Table 5.91 provides the N-Values by Time Point.

Group Baseline Week 2 Week 6 Week 10 Pre-Necropsy 1 8 7 5 4 3 2 8 8 6 34 3 8 7 7 3 4 4 8 7 4 2 2

Ultrasonography on the kidneys over the course of the study showedincreases in surface area (length×width) indicating kidney changes(hydronephrosis) in all treatment groups. Group averages were similarover time for all groups for the right and left kidney (Table5.92—Average Kidney Surface Area by Group for 12 non-PCV-2 UnscheduledDeaths (L×W, cm2).

TABLE 5.92 Group Baseline Week 2 Week 6 Week 10 Pre-Necropsy Left Kidney1 (N = 1) 21.0 NA NA NA NA 2 (N = 3) 14.5 19.8 15.5 13.1 18.8 3 (N = 4)19.5 20.9 22.3 14.4 42.5 4 (N = 4) 13.9 21.7 20.2 NA NA Right Kidney 1(N = 1)* 21.2 NA NA NA NA 2 (N = 3) 12.9 16.4 18.7 12.3 20.0 3 (N = 4)19.7 19.0 30.6 13.9 14.7 4 (N = 4) 15.5 14.4 19.2 NA NA NA = Notapplicable *Animal no. 8 euthanized on day 9.

Ultrasonography on the implant showed a decrease in wall thickness overtime in all of the construct groups (Groups 1-3). The scaffold onlygroup (Group 4) showed a slight increase in thickness; however there arelimited data points for analysis (Table 5.93—Average Conduit WallThickness by Group for 12 non-PCV-2 Unscheduled Deaths (cm).

TABLE 5.93 Pre- Group Week 2 Week 6 Week 10 Necropsy 1 (N = 1)* NA NA NANA 2 (N = 3) 0.225 0.201 0.157 0.131 3 (N = 4) 0.291 0.184 0.228 0.144 4(N = 4) 0.203 0.358 NA NA NA = Not applicable *Animal no. 8 euthanizedon day 9.

Pathology. The pathology report appears in the Example below.

Viral infection and unscheduled deaths from consequences of obstructionreduced the number of animals surviving for study duration from 32 to atotal of 7. For the animals surviving to sacrifice (N=7), adhesions werepresent in all animals and enteric-conduit fistulas were present in 4 of7 animals (Pathology Example below). Hydroureter and hydronephrosis(unilateral or bilateral) were present in all animals, andpyelonephritis was present in 1 of 7 animals. The animal withpyelonephritis was in the scaffold-only treatment group (Group 4). Forthe non-PCV-2 unscheduled deaths (N=12), adhesions were present in allanimals and fistulas were present in 7 of 12 animals (enteric-conduit)and 1 of 12 animals (ureter-intestine). Hydroureter and hydronephrosis(unilateral or bilateral) were present in 8 of 12 animals, andpyelonephritis was present in 4 of 12 animals. Three animals withpyelonephritis were in the scaffold only treatment group (Group 4), and1 animal was in the blood-derived autologous SMC treatment group (Group3). Despite complications that led to unscheduled deaths, the constructimplants served as templates for a regenerative process that resulted ina tissue comprised of urothelium and smooth muscle layers. Regenerationwas most prominent at the ureteral end of the implant. The tissue in thecaudal implant and atrium segments of the conduit had a wall comprisedof fibrous connective tissue without urothelial lining The regenerativeprocess at the ureteral end of the implant resulted in urinary tissueformation that was comparable among construct groups and distinct fromthe reparative healing which typified the scaffold-only group.

The scaffold-only test article was determined to be unsuitable forfurther development because the outcome was not consistent with normalurinary tissue and these animals had a higher incidence of bilateralrenal complications. Outcomes observed in construct groups, regardlessof cell source, were equivalent.

DISCUSSION. Of the 31 animals recovered from surgery, seven animals(23%; two from each of Groups 1, 3 and 4; and one from Group 2)successfully completed the full duration of the in-life portion of thestudy. Macroscopic pathology was evident in all groups and includedevidence of viral infection, intermittent obstruction of urine flow,debris and detritus buildup in the conduit, abdominal and pelvicadhesions, fistulae, hydronephrosis, and hydroureter and pyelonephritis.Evidence of urinary tissue regeneration was observed to varying degreesin construct implants seeded with SMC from each source (Groups 1-3,bladder, adipose, and blood derived SMC, respectively). However,incomplete urinary tissue healing and increased incidence of upperurinary tract pathology were observed in the scaffold-only implantedanimals (Group 4). Twelve of the 31 animals (39%) were infected withPCV-2 virus. Although a viral infection resulted in reduced survival,urinary tissue regeneration was also observed in the construct groups(Groups 1-3). The surgical placement of the test articles was designedfor animal welfare purposes to achieve optimal voiding of urine byquadrupeds so as to avoid irritant contact dermatitis from urine andpotential early morbidity and euthanasia. However, this surgicalapproach resulted in test articles being directly under the overlyingabdominal organs and the weight of these organs and the pressure of theabdominal viscera caused intermittent conduit obstruction. Thisquadruped specific surgical placement may have also contributed to theobserved adhesions, fistulae, and upper urinary tract complications(e.g., dilation, inflammation, and/or infection of ureters or kidney).However, intestinal obstruction was not observed in this study. The pigis commonly used as an animal model for studying the development andprevention of post-surgical adhesions therefore some incidence ofadhesions was expected simply from the abdominal laparotomy for biopsycollection in construct animals (Groups 1-3). Animals in Groups 1-3 alsounderwent a second surgical procedure to implant the test article,substantially increasing the risk of adhesions compared to the animalsreceiving scaffold-only implants that underwent a single surgicalprocedure (test article implantation only). Adhesions observed in thisstudy were considered a consequence of these surgical procedure(s).

All the adverse findings noted in the upper urinary tract (hydroureter,hydronephrosis, and pyelonephritis) were consequences of conditions thatintermittently obstructed the flow of urine through the conduit. Thepyelonephritis observed in the quadruped pigs of this study wasconsidered secondary to the debris and detritus build-up and bacterialcontamination of the stoma from feces and skin. Pyelonephritis was mostfrequently observed in the scaffold-only animals (Group 4). The surgicalpositioning of the test article for all animals and multiple surgeriesfor animals in Groups 1-3 (biopsy and test article implantation)promoted the formation of abdominal and pelvic adhesions. Furthermore,the intermittent obstruction, the debridement protocol used for stomamanagement (use of forceps in tranquilized animals to remove detritusbuildup), viral infection, and adhesions of intestinal tract to theconduit contributed to enteric-conduit fistula formation and upper renaldisease. Despite complications that led to unscheduled deaths, theconstruct implants (Groups 1-3) served as templates for a regenerativeprocess that resulted in a tissue comprised of urothelium and smoothmuscle layers (native urinary tissue components). Regeneration was mostprominent at the ureteral end of the implant. The tissue in the atrium(peritoneal tissue only with no construct or scaffold) segments of theconduit had a wall comprised of fibrous connective tissue withouturothelial lining, indicating the peritoneum alone was incapable ofsupporting regeneration of urinary tissue. The regenerative process atthe ureteral end of the implant resulted in urinary tissue formationthat was comparable among construct groups and distinct from thereparative healing which typified the scaffold-only group. Thescaffold-only test article outcome was not consistent with normalurinary tissue and the wall was comprised of fibrovascular connectivetissue that could be predisposed to scarring and/or stenosis of thelumen. The incomplete formation of urinary tissue in the animalsreceiving a scaffold-only implant exacerbated the upper urinary tractfindings from intermittent obstruction discussed above; Group 4 animalshad a higher incidence of the bilateral renal complications. Outcomesobserved in construct groups, regardless of cell source, wereequivalent.

CONCLUSION. The objective of this study was to evaluate the safety andfunctionality of a Neo-Urinary Conduit (NUC) seeded with autologousbladder-derived, adipose-derived or blood-derived smooth muscle cells(SMC), or scaffold-only (scaffold without SMC). Functionalityassessments were urinary flow and urinary tissue regeneration. Only 7 of31 (23%) animals completed this study. The primary safety findingsunderlying the 24 of 31 unscheduled deaths were PCV-2 viral infectionand intermittent obstruction of the conduit resulting in upper urinarytract injury (hydronephrosis, hydroureter, and pyelonephritis).Obstruction was managed by debridement and saline flushing of the stoma.Urinary tissue-like regeneration characterized by mucosa, submucosa andsmooth muscle with a fibrovascular stroma was observed after constructtest article implantation regardless of SMC source (i.e. bladder,adipose, or blood). In contrast, a reparative process was observedfollowing implantation of the scaffold-only test article characterizedby an abnormal mucosa supported by fibrovascular stroma with limitedsmooth muscle. The predisposition of the pig to adhesion formation andthe two surgical procedures related to tissue biopsy and test articleimplantation in all construct groups resulted in adhesions between theregenerated tissue and overlying bowel. Fistula formation wasexacerbated by frequent stoma cleaning and the pig's quadruped stance.The absence of SMC in the scaffold-only test articles appears to havecontributed to incomplete development of urinary tissue and an increasein mortality subsequent to obstruction, leading to the determinationthat the scaffold-only test article was unsuitable for furtherdevelopment.

Example 6 Pathology of Animals Following Implantation of Neo-UrinaryConduit Constructs

At the conclusion of the study described in Example 5, the anatomicpathology of the test animals was assessed.

Tissue Collection. As per protocol, the abdominal cavity was opened andthe conduit (the outcome of implanting a construct or scaffold-only testarticle), was visualized and digitally photographed in situ at theanimal facility. The conduit was removed en bloc with the kidneys andureters. The ureters were measured, and then detached from the conduitby transverse sectioning 3-4 cm away from the anastomoses.Representative sections of the kidneys, ureters, lymph nodes, and anyother lesions observed grossly were obtained. All tissue samples wereplaced in 10% Neutral Buffered Formalin (NBF) for histologicalprocessing and evaluation.

Histological Processing. After fixation, the conduit was openedlongitudinally (parallel with the outflow) and divided into dorsal andventral halves as illustrated in the trimming diagram below.

FIG. 59A-B shows a post-fixed conduit tissue from DB-252 Animal no. 18and a trimming schematic. FIG. 59C shows the position of the ureters,stoma, mid-conduit, and a diverticulum.

Three transverse sections were trimmed from the body of each half(cranial, mid and caudal captured on slides 5, 6, 7 on the dorsal halfand 9, 10, 11 on the ventral half, respectively). One section from eachhalf was taken from the atrium (captured on slide 8 on the dorsal halfand 12 on the ventral half). An additional section was taken at each ofthe two ureter-conduit junctions (left captured on slide 13, right onslide 15). One other slide (number 17) was used to capture the stoma atthe skin surface and the adjacent canal through the abdominal wall. Whenthe size of the conduit permitted, this scheme resulted in 11 slides.Some conduits were too short in length to accommodate the trimmingscheme, so the available conduit was divided into fewer sections. Thesections collected from each animal include conduit dorsal cranial,conduit dorsal mid, conduit dorsal caudal, dorsal stomal conduitjunction, conduit ventral cranial, conduit ventral mid, ventral conduitstomal junction, left ureter conduit stomal junction, rightureter-conduit junction, and conduit stoma-skin junction. In addition,the following tissue/organ sections were obtained and submitted forhistology: left kidney (slide 1), right kidney (slide 2), left ureter(slide 14), right ureter (slide 16), lumbar lymph node (slide 3),mesenteric lymph nodes (slide 4) and any gross lesions (slides 18, 19etc.). During trimming of tissues, digital photographs were taken forillustration purposes. Post fixation, tissues were processed routinelyto microslides and stained with hematoxylin and eosin (H&E) and Masson'strichrome. Slides were evaluated microscopically. Where appropriate,microscopic observations for individual animal data were obtained andscored.

Results

Mortality. During definitive surgery, the ureters of Animal no. 30(Group 4) were perforated, thus preventing test article implantation.The animal was euthanized and not replaced, reducing the total N ofanimals implanted with test articles (construct or scaffold-only) to 31and the N of Group 4 to 7 animals. Twenty-four animals were unscheduleddeaths and seven animals survived until the scheduled sacrifice.Survival days post-implant and final disposition of each of the 31animals implanted with the test article are shown in Appendix 1, AnimalInformation. Disposition (mortality classification) of the 24unscheduled deaths is summarized by treatment group in Table6.2—Unscheduled deaths by Group)

TABLE 6.2 Mortality Group 1 Group 2 Group 3 Group 4 Total Group N 8 8 87 31 In-life procedure-related 0/8 0/8 1/8 0/7 1/31 death Found dead 0/82/8 0/8 1/7 3/31 Euthanized 6/8 5/8 5/8 4/7 20/31  Total Unscheduled 6/87/8 6/8 5/7 24/31  Necropsies Scheduled Necropsies 2/8 1/8 2/8 2/7 7/31

The following sections discuss the underlying findings observed forunscheduled deaths.

Viral Related Mortalities. During in-life clinical observations, skinlesions consistent with PCV-2 infection were observed. Twelveunscheduled death animals developed one or more lesions, consistent withdistinct pathological features ascribed to porcine dermatitis andnephropathy syndrome (PDNS), associated with PCV-2 infection. Animalswere classified as PCV-2-infected if any of the following were observed:

-   1) Clinically-observed purple skin discoloration-   2) Microscopic vasculitis or vasculitis/perivasculitis affecting the    kidney, skin or lung-   3) Kidney findings of tubular    necrosis/fluid/casts/glomerulonephritis, or viral inclusions of    tubular epithelial cells-   4) Lymphocyte depletion in lumbar lymph nodes in the presence of 1,    2, or 3

By these criteria, porcine PCV-2 infection was identified in 12 of the24 unscheduled necropsy animals. These included 5 animals in Group 1(animal nos. 5113342, 7, 2, 4 and 3); 4 animals in Group 2 (animal nos.13, 16, 14 and 10); 2 animals in Group 3 (animal nos. 20 and 19); and 1animal in Group 4 (animal no. 28). The microscopic findings from thePCV-2 animals will not be discussed in the results section as the PCV-2condition was deemed an assignable cause of morbidity unrelated to thedevice.

Non-PCV-2 Associated Mortalities. A total of 12 unscheduled deaths werenot associated with a detected PCV-2 virus infection (including the 1in-life procedure-related death). The 12 unscheduled deaths notattributable to PCV-2 infection included 1 animal in Group 1 (animal no.8); 3 animals in Group 2 (animal nos. 11, 9, and 12), 4 animals in Group3 (animal nos. 24, 21, 22, and 17) and 4 animals in Group 4 (animal nos.32, 31, 29, and 27). An important event in the clinical decline of theseanimals was obstruction of the outflow of urine and debris through thestoma. Obstruction appeared to have been facilitated by the placement ofthe test article in the ventral portion of the abdominal cavity wherethe weight of the overlying abdominal organs could lead to conduitclosure, adhesion and fistula formation, and renal complications (e.g.,dilation, inflammation, and/or infection of ureters or kidney). Surgicalplacement of the test article was the same in all study animals;therefore, obstruction-related complications had a similarpathobiological mechanism (i.e. abdominal viscera resting on the conduitbecause of the quadruped anatomy) in all groups.

Distribution of Unscheduled Deaths by Underlying Findings. A summary ofunderlying findings in the 24 unscheduled deaths is presented in Table6.3—PCV-2 and Non-PCV-2 Associated Unscheduled Deaths by Group.

TABLE 6.3 Mortality Categories Group 1 Group 2 Group 3 Group 4 TotalEvidence of PCV-2 5 4 2 1 12 infection Non-PCV-2 associated 1 3 4 4 12unscheduled deaths Total unscheduled deaths 6 7 6 5 24

Findings Pertinent to Safety. A comprehensive list of macroscopicfindings and microscopic correlates for all animals was prepared (datanot shown). Individual microscopic animal data was obtained (data notshown). Discussion of macroscopic and microscopic findings is focused tothose relevant to the obstruction of urine outflow that was facilitatedby the anatomical placement of test articles in the 7 animals survivingto scheduled sacrifice and the 12 non-PCV-2 associated unscheduleddeaths (N=19). Conduits formed from implantation of a test article werevariably sized and shaped tubes located in the retro-peritoneal space ofthe ventral abdomen. The ureters entered at the cranial end of theconduit (ureter-conduit junction, UCJ, FIG. 59). Urine flow was directedthrough the peritoneal wrapped implant and the atrium and emerged at thestoma. The cranial end of the conduit frequently had bilateral bulbousdilations, referred to as diverticula that were considered to be theconsequence of intermittent obstruction.

Adhesions and Fistulas. The ventral side of the conduit was adhered tothe fascia and skeletal muscle of the abdominal wall, and the dorsalside was covered with peritoneum. At necropsy, the conduit was oftendifficult to visualize because of marked adhesions (e.g., conduit togastrointestinal tract, omentum, uterus, seminal vesicle, liver,pancreas, spleen, or reproductive organs). The lumen of the conduit wasoften filled with detritus. Fistulas were observed between conduit andadjacent hollow organs (e.g., intestine). Specifically,conduit-intestine or conduit-ureter fistulas were observed in the 19animals evaluated.

Adhesions and Fistulas in 7 Surviving Animals. Adhesions were present inall 7 animals surviving to the scheduled sacrifice (Table 6.4). Fistulasbetween conduit and intestinal tract were observed in 4 of these 7animals: 2/2 in Group 1 (animal nos. 1 and 6); 1/1 in Group 2 (animalno. 15) and 1/2 in Group 3 (animal no. 23). Table 6.4: Incidence ofAdhesions and Fistulas in 7 Surviving Animals.

TABLE 6.4 Group 1 Group 2 Group 3 Group 4 Macroscopic Finding N = 2 N =1 N = 2 N = 2 Total Adhesions 2/2 1/1 2/2 2/2 7/7 Conduit-IntestineFistula 2/2 1/1 1/2 0/2 4/7

Adhesions and Fistulas in 12 Unscheduled Deaths. Adhesions were presentin all 12 non-PCV-2 associated early deaths (Table 6.5). The presence offistulas between conduit and intestinal tract was observed in 7 of 12animals: 3/3 in Group 2 (animal nos. 11, 12 and 9), 3/4 in Group 3(animal nos. 21, 22, and 17) and 1/4 in Group 4 (Animal No. 29). Onefistula between intestine and ureter was observed in a Group 4 animal(animal no. 27). Table 6.5: Incidence of Adhesions and Fistulas in 12non-PCV-2 Unscheduled Deaths.

TABLE 6.5 Group 1 Group 2 Group 3 Group 4 Macroscopic Finding N = 1 N =3 N = 4 N = 4 Total Adhesions 1/1 3/3 4/4 4/4 12/12  Conduit-IntestineFistula 0/1 3/3 3/4 1/4 7/12 Ureter-Intestine Fistula 0/1 0/3 0/4 1/41/12

Ureters and Kidneys. Thickened ureters observed macroscopically resultedfrom several underlying phenomena upon microscopic evaluation. Ureterdilatation (or hydroureter) was characterized by an expanded lumen withnormal ureteral wall structure. Transitional cell hyperplasia wascharacterized by increased number of cells in the transitionalepithelium, and vacuolation was characterized by round, clear vacuolesin the epithelium. Subacute/chronic inflammation of the peri-uretermesentery occurred when the mesentery surrounding the ureter wasexpanded by collagen and fibroblasts, with occasional lymphocytes andmacrophages. This inflammation did not usually affect the muscle tunicsor urothelium of the ureter. Peri-ureter inflammation could be relatedto adhesion between ureter and intestinal or reproductive organs; but itcould also occur without adhesion.

Microscopically, hydronephrosis was characterized by a dilatation of therenal pelvis with thinning and chronic inflammation (fibrosis,lymphocytes, plasma cells and occasional macrophages) of the renalcortex. Hydronephrosis was considered to be the result of full orpartial obstruction in the lower urinary system (ureters, conduit oratrium/stoma). Chronic-active pyelonephritis, which was often associatedwith hydronephrosis, was characterized by infiltration of neutrophilsand cellular debris into the renal pelvis, often spreading into thedistal medulla.

Pyelonephritis was considered to be the result of bacterial infection ofthe lower urinary tract which ascended into the renal pelvis. Chronicnephritis (without hydronephrosis) was characterized by fibrosis withinfiltration of inflammatory cells (lymphocytes, macrophages, plasmacells and occasionally neutrophils) in the renal cortex or medulla.

The cortex of kidneys with chronic nephritis looked similar to those inanimals with hydronephrosis/chronic nephritis; however, in chronicnephritis the pelvis was not dilated. Chronic-active nephritis wassimilar in appearance to chronic nephritis, but with significantinfiltration of neutrophils. Tubularnecrosis/fluid/casts/glomerulonephritis was a constellation of changescharacterized by neutrophils, lymphocytes and macrophages in glomeruli,necrosis of individual tubular epithelial cells, proteinaceous tubularcasts and/or hemorrhage in tubular lumens.

Hydroureter and Hydronephrosis and/or Pyelonephritis in 7 SurvivingAnimals. Hydroureter and hydronephrosis were observed microscopically inall 7 surviving animals (Table 6.6). Unilateral hydroureter (2/7animals): 2 animals in Group 3 (animal nos. 23 and 18). Bilateralhydroureter (5/7 animals): 2 animals in Group 1 (animal nos. 1 and 6), 1animal in Group 2 (animal no. 15), and 2 animals in Group 4 (animal nos.25 and 26). Unilateral hydronephrosis (6/7 animals): 2 animals in Group1 (animal nos. 1 and 6), 1 animal in Group 2 (animal no. 15), 2 animalsin Group 3 (animal nos. 18 and 23), and 1 animal in Group 4 (animal no.26). Bilateral hydronephrosis (1/7 animals); Group 4 (animal no. 25).Pyelonephritis (unilateral) was observed in 1 of 7 animals; Group 4animal no. 26. Table 6.6: Incidence of Hydroureter, Hydronephrosisand/or Pyelonephritis in 7 Surviving Animals.

TABLE 6.6 Group 1 Group 2 Group 3 Group 4 Finding N = 2 N = 1 N = 2 N =2 Total Hydroureter, Unilateral 0/2 0/1 2/2 0/2 2/7 Hydroureter,Bilateral 2/2 1/1 0/2 2/2 5/7 Hydronephrosis, Unilateral 2/2 1/1 2/2 1/26/7 Hydronephrosis, Bilateral 0/2 0/1 0/2 1/2 1/7 Pyelonephritis,Unilateral 0/2 0/1 0/2 1/2 1/7 Pyelonephritis, Bilateral 0/2 0/1 0/2 0/20/7

Hydroureter and Hydronephrosis and/or Pyelonephritis in 12 UnscheduledDeaths. Hydroureter and/or hydronephrosis were observed microscopicallyin 8/12 and 8/12 of the non-PCV-2 associated early deaths, respectively(Table 6.7). Unilateral hydroureter (3/12 animals): 1 animal in Group 2(animal no. 12), 1 animal in Group 3 (animal no. 21) and 1 animal inGroup 4 (animal no. 27). Bilateral hydroureter (5/12 animals): 2 animalsin Group 3 (animal nos. 22 and 24), and 3 animals in Group 4 (animalnos. 32, 31 and 29). Unilateral hydronephrosis (5/12 animals): 2 animalsin Group 2 (animal nos. 9 and 12), 1 animal in Group 3 (animal no. 21),and 2 animals in Group 4 (animal nos. 32 and 27). Bilateralhydronephrosis (3/12 animals): 1 animal in Group 3 (animal no. 24) and 2animals in Group 4 (animal nos. 31, and 29).

Unilateral pyelonephritis was observed in 2 of 12 animals): 1 animal inGroup 3 (animal no. 24) and 1 animal in Group 4 (animal no. 32).Bilateral pyelonephritis was observed in 2 of 12 animals): 2 animals inGroup 4 (animal nos. 31 and 29). Table 6.7: Incidence of Hydroureter,Hydronephrosis and/or Pyelonephritis in 12 non-PCV-2 Unscheduled Deaths

TABLE 6.7 Group 1 Group 2 Group 3 Group 4 Finding N = 1 N = 3 N = 4 N =4 Total Hydroureter, Unilateral 0/1 1/3 1/4 1/4 3/12 Hydroureter,Bilateral 0/1 0/3 2/4 3/4 5/12 Hydronephrosis, Unilateral 0/1 2/3 1/42/4 5/12 Hydronephrosis, Bilateral 0/1 0/3 1/4 2/4 3/12 Pyelonephritis,Unilateral 0/1 0/3 1/4 1/4 2/12 Pyelonephritis, Bilateral 0/1 0/3 0/42/4 2/12

Findings Pertinent to Regeneration of a Urinary Conduit. Tissuecomponents from each section were observed (data not shown). Thediscussion of findings pertinent to regeneration below focuses on thesections obtained for histological evaluation (FIG. 59) from the 7animals surviving to scheduled sacrifice and the 12 unscheduled deaths.The conduit that developed after surgical implantation of the testarticle consisted of a central lumen that coursed from ureters (cranialend) through the implant and atrium to the stomal opening in the skin ofthe ventral abdomen. The histological appearance of the conduit wallvaried depending upon location of sample within the conduit and animalsurvival time.

Ureter-Conduit Junction and Cranial Portion of Conduit. The typicalcomposition of tissue near the cranial end of the conduit encompassingthe ureterconduit junction (UCJ; Sections 13 and 15 in FIG. 59) and thecranial portion (Sections 5 and 9 in FIG. 59) was urothelium overlyinglayers of smooth muscle fibers with interspersed connective tissue. UCJand cranial sections of the conduit were morphologically similar to theureters, although the conduit wall thickness was typically greater thanthat of the ureter, particularly within diverticula. Diverticulaappeared as bi-compartmental portions of conduit projecting caudallyfrom the left and right ureteral-conduit junctions. The typicalappearance of urothelium was minimally-to-moderately vacuolated, andvariable in thickness. Urothelium thickness varied betweenminimally-to-moderately attenuated (especially within a largediverticulum) and minimally-to-mildly hyperplastic. For the 7 animalssurviving to scheduled sacrifice, incidence of urothelium and smoothmuscle layers observed in Sections 5, 9, 13, and 15 was similar amongall animals. Table 6.8: Incidence of Urothelium and Smooth Muscle inCranial Conduit in 7 Surviving Animals

TABLE 6.8 Group 1 Group 2 Group 3 Group 4 Histological Finding N = 1 N =2 N = 2 N = 2 Presence of Urothelium UCJ (slides 13 or 15) 2/2 1/1 2/22/2 Cranial Conduit (slides 5 or 9)  1/2* 1/1  1/2* 1/2 Presence ofSmooth Muscle UCJ (slides 13 or 15) 2/2 1/1 2/2 2/2 Cranial Conduit(slides 5 or 9)  1/2* 1/1  1/2* 1/2 *Due to size of regenerated conduit,insufficient tissue available to evaluate section(s) in 1 animal pergroup in Group 1 & 3.

For the 12 non-PCV-2 associated early deaths, the incidence ofurothelium and smooth muscle layers in Sections 5, 9, 13, and 15 wassimilar among animals within and between groups receiving constructimplants (Groups 1-3). The incidence of urothelium and smooth musclelayers in Sections 5, 9, 13, and 15 among animals receivingscaffold-only implants (Group 4) was lower than that observed in animalsreceiving a construct implant. Table 6.9: Incidence of Urothelium andSmooth Muscle in Cranial Conduit in 12 Unscheduled Deaths.

TABLE 6.9 Group 1* Group 2 Group 3 Group 4 Histological Finding N = 1 N= 3 N = 4 N = 4 Presence of Urothelium UCJ (slides 13 or 15) 0/1 3/3 4/43/4 Cranial Conduit (slides 5 or 9) 0/1 2/3 2/4{circumflex over ( )} 0/4Presence of Smooth Muscle UCJ (slides 13 or 15) 0/1 2/3 4/4 2/4 CranialConduit (slides 5 or 9) 0/1 2/3 2/4{circumflex over ( )} 0/4 *animal onstudy 9 days {circumflex over ( )}Due to size of regenerated conduit,insufficient tissue available to evaluate section(s) in one animal inGroup 3

Mid and Caudal Portions of Conduit. For all 19 animals, the typicalappearance of the mid (Sections 6 and 10 in FIG. 59) and caudal(Sections 7 and 11 in FIG. 59) portions of the conduit differed fromthat observed in the UCJ and cranial sections. The point of transitionfrom conduit to atrium varied between animals because the caudal end ofthe implant floated freely within the peritoneal wrapping making thetransition from conduit to atrium difficult to define at necropsy. Thetypical composition of Sections 6, 7, 10, and 11 was organized collagenwith associated fibroblasts. Internal to the collagenous wall andclosest to the lumen was a layer of chronic-active inflammationcomprised of loosely arranged collagen, capillaries and abundantneutrophils with fewer lymphocytes and macrophages. Internal to theinflammation, the lumen was often filled with detritus comprised ofdegenerate or necrotic inflammatory cells (primarily neutrophils) andcellular debris with admixed bacterial colonies.

Atrium and Stoma Portions of Conduit. At the region of the atrium-stomalend of the conduit (Sections 8, 12 and 17 in FIG. 59), the stoma-atriumjunction was visible where the organized collagen and adnexa of thestomal dermis apposed the collagenous wall (without adnexa). For all 19animals, these sections consisted mainly of squamous epithelium andchronic-active inflammation/detritus. Typically, the squamous epithelium(epidermis) of the skin extended cranially for a short distance over theatrium. The external surface of the atrium was comprised of looseconnective tissue of peritoneum origin. This external covering was theequivalent of a serosal layer and contained nerves, blood vessels,adipose tissue, and some areas of fibrous connective tissue (collagenfibers and fibroblasts). At the site of one of the fistulas, ectopicintestinal mucosa was observed to cover a small segment of conduitadjacent to fistulas. This was noted in one of the 7 surviving Group 2animals (animal no. 15).

Healing Outcomes in Animals Receiving Construct or Scaffold-onlyImplants. Urinary tissue-like regeneration characterized by mucosa,submucosa and smooth muscle with a fibrovascular stroma was observedafter construct test article implantation regardless of SMC source (i.e.bladder, adipose or blood). Areas of bladder-like tissue, comprised ofcontinuous urothelium with underlying smooth muscle, were observed in 4of 5 surviving animals and 4 of 8 unscheduled sacrifice animalsimplanted with construct test articles. In the majority of scaffold onlyanimals, the conduit tissue morphology was compatible with a reparativeprocess, characterized by urothelium extending only a short distance(approximately 1 mm) from the ureter-conduit junction without underlyingsmooth muscle.

Discussion. Seven of 31 animals survived to scheduled necropsy and overthe course of the study 24/31 animals were euthanized off-schedule orfound dead. Viral infections contributed to 12/24 unscheduled deaths.There were both clinical and pathological signs compatible withconcurrently infection with PCV-2 during the course of this study.Macroscopic findings were ecchymoses (clinically, purple discoloration)of the skin and enlarged inguinal lymph nodes (macroscopic finding ofviral infections were observed in 11/24 unscheduled deaths). Microscopicfindings included renal changes consisting of tubularnecrosis/fluid/casts/glomerulonephritis, eosinophilic intracellularinclusions in tubular epithelial cells in the kidney, andvasculitis/perivascular inflammation in the kidneys and ureters, andinflammation in the lungs (microscopic findings revealed an additionalanimal with PCV-2 infection at necropsy, bringing the total viralinfection related unscheduled deaths to 12/24). These findings are amongthose commonly reported for pigs with PCV-2 infection. The infectionwith PCV-2 contributed to the poor clinical health of the animals (e.g.lethargy, diarrhea, loss of appetite) and subsequent humane earlyeuthanasia. Twelve of 24 unscheduled deaths were non-PCV-2 associated:1/24 unscheduled death was the result of an in-life proceduralcomplication and obstruction contributed to 11/24 unscheduled deaths.Obstruction was caused by a combination of the surgical placement of thetest article along the abdominal floor of the pig where the weight ofabdominal viscera compressed the implant and the use of peritoneum toform an atrium segment connecting the conduit to the skin resulting indetritus buildup from the external environment and mucus in the urine(normal for swine).

Relevant postoperative complications that are inherent in this quadrupedanimal model include: (i) abdominal adhesions leading to potentialfistula formation and (ii) location of the test article placement inrelation to the abdominal organs.

When intestines were the adhered organ, the tunica muscularis of theadhered segment of intestine was often diminished or eroded at the pointof adhesion, as was the atrium wall. Test articles were placed in theintra-abdominal cavity of the ventral abdomen with a peritoneum-derivedtube extending from the caudal end of the test article through theabdominal wall to the skin. The test article was anchored at the cranialend to the ureters, but floated freely within the peritoneal wrapping atthe caudal end. Healing resulted in a urinary conduit consisting of acentral lumen that coursed from ureters (cranial end) to the stomalopening in the skin of the ventral abdomen (caudal end). Urothelium andlayered smooth muscle formation were most frequently observed at thecranial end of the conduit where the implanted test article was presentwithin the peritoneal wrap. The atrium formed from peritoneum only wascomprised of fibrous connective tissue walls without urothelialcovering.

The incidence of urothelium and/or smooth muscle layers in the cranialconduit (near the ureters) was higher in the construct groups (Groups 1,2 and 3) than the scaffold-only group (Group 4). The extent ofregeneration observed in sections taken near the caudal end of theimplant was variable because the boundary between the caudal end of theimplant and the cranial side of the atrium was difficult to discern atnecropsy. Most sections presumed to be at the midsection of the implant(Sections 6 and 10, FIG. 59) had no urothelium present. Table 6.10 showsa summary of the findings by group for the neo-urinary conduit.

TABLE 6.10 Group Treatment 1 2 3 4 Autologous Bladder AutologousAutologous Scaffold SMC's Adipose SMC's Blood SMC's Only Mean Days onStudy 53 60 61 48 Incidence of Select Findings by Conduit Location (#animals with finding/# animals examined at that location) Presence ofUrothelium Left Ureter Conduit Junction (Slide13) 5/8 6/8 7/8 3/7 RightUreter Conduit Junction (Slide 15) 7/8 6/8 4/7 4/7 Ureter End of ConduitBody (Slides 5 or 9) 3/7 5/8 3/5 1/7 Middle of Conduit Body (Slides 6 or10) 0/6 1/7 1/5 1/7 Stoma End of Conduit Body (Slides 7, 8, 11 0/8 0/80/7 0/7 or 12) Conduit-Stoma-Skin Junction (Slide 17) 1/8 0/8 0/8 0/7Presence of Squamous Epithelium Left Ureter Conduit Junction (Slide 13)0/8 0/8 0/8 0/7 Right Ureter Conduit Junction (Slide 15) 0/8 0/8 0/8 0/7Ureter End of Conduit Body (Slides 5 or 9) 0/7 0/8 1/5 0/7 Middle ofConduit Body (Slides 6 or 10) 0/6 1/7 0/5 0/7 Stoma End of Conduit Body(Slides 7, 8, 11 6/8 3/8 4/7 2/7 or 12) Conduit-Stoma-Skin Junction(Slide 17) 7/8 8/8 8/8 7/7 Presence of Smooth Muscle Left Ureter ConduitJunction (Slide13) 4/8 4/8 7/8 2/7 Right Ureter Conduit Junction (Slide15) 4/8 4/8 5/8 3/7 Ureter End of Conduit Body (Slides 5 or 9) 3/7 5/83/5 1/7 Middle of Conduit Body (Slides 6 or 10) 0/6 1/7 1/5 1/7 StomaEnd of Conduit Body (Slides 7, 8, 11 0/8 0/8 0/7 0/7 or 12)Conduit-Stoma-Skin Junction (Slide 17) 0/8 0/8 0/8 0/7 Surface ofchronic-active inflammation/detritus Left Ureter Conduit Junction(Slide13) 5/8 5/8 1/8 6/7 Right Ureter Conduit Junction (Slide 15) 3/85/8 3/8 5/7 Ureter End of Conduit Body (Slides 5 or 9) 5/7 6/8 3/5 6/7Middle of Conduit Body (Slides 6 or 10) 6/6 7/7 5/5 6/7 Stoma End ofConduit Body (Slides 7, 8, 11 8/8 7/8 7/7 7/7 or 12) Conduit-Stoma-SkinJunction (Slide 17) 6/8 4/8 5/8 6/7 Attenuation, Urothelium 1/8 3/8 3/80/7 Hyperplasia, Urothelium 3/8 6/8 5/8 4/7 Vacuolation, Urothelium 6/86/8 6/8 4/7 Ectopic GI mucosa 1/8 2/8 0/8 0/7 Heterotopic bone,sub-urothelial 1/8 0/8 2/8 0/7 Hemorrhage, conduit wall 0/8 3/8 1/8 2/7Mineralization 1/8 0/8 0/8 0/7 Scaffold material 1/8 0/8 1/8 1/7 AdheredGI tract (P = present) 6/8 6/8 1/8 5/7 Adhered spleen (P = present) 0/80/8 0/8 1/7 Adhered seminal vesicle/uterus 0/8 0/8 2/8 0/7 (P = present)Vasculitis/vascular necrosis, small blood 3/8 0/8 2/8 1/7 vessels Acuteinflammation/ulceration, skin near 1/8 0/8 0/8 0/7 stoma

Table 6.11 shows a summary of the findings by group for kidney, ureterand other tissues.

TABLE 6.11 Group Treatment 1 2 3 4 Autologous Bladder AutologousAutologous Scaffold SMC's Adipose SMC's Blood SMC's Only Left KidneyHydronephrosis/chronic nephritis 3/8 2/8 2/8 6/7 Chronic nephritis(without 1/8 2/8 1/8 0/7 hydronephrosis) Pyelonephritis 2/8 0/8 0/8 3/7Chronic-active nephritis 1/8 4/8 1/8 5/7 Inflammation, acute/subacute1/8 1/8 0/8 0/7 Regeneration, tubular epithelium 5/8 6/8 4/8 1/7 Tubular1/8 2/8 0/8 1/7 necrosis/fluid/casts/glomerulonephritis Viralinclusions, tubular epithelial 1/8 2/8 0/8 0/7 cellsVasculitis/perivascular inflammation 1/8 1/8 1/8 0/7 Inflammation,subacute, pelvis 2/8 0/8 1/8 0/7 Hyperplasia, transitional epithelium,1/8 0/8 1/8 0/7 pelvis Inflammation, chronic-active, 0/8 2/8 2/8 0/7capsule/peritoneum Right Kidney Hydronephrosis/chronic nephritis 3/8 2/83/8 5/7 Chronic nephritis (without 2/8 1/8 2/8 0/7 hydronephrosis)Pyelonephritis 0/8 0/8 1/8 3/7 Chronic-active nephritis 1/8 2/8 0/8 1/7Inflammation, acute/subacute 1/8 0/8 0/8 0/7 Regeneration, tubularepithelium 4/8 2/8 4/8 1/7 Tubular 1/8 2/8 0/8 0/7necrosis/fluid/casts/glomerulonephritis Viral inclusions, tubularepithelial 1/8 1/8 0/8 0/7 cells Vasculitis/perivascular inflammation3/8 0/8 2/8 0/7 Inflammation, subacute, pelvis 1/8 0/8 0/8 0/7Hyperplasia, transitional epithelium, 1/8 0/8 1/8 1/7 pelvisInflammation, chronic-active, 1/8 2/8 0/8 0/7 capsule/peritoneum LeftUreter Dilatation 4/8 2/8 3/8 7/7 Hyperplasia, transitional epithelium1/8 0/8 0/8 0/7 Vacuolation, transitional epithelium 1/8 0/8 1/8 0/7Inflammation, subacute/chronic, peri- 3/8 4/8 5/8 3/7 ureter mesenteryVasculitis/necrosis, mesenteric blood 4/8 1/8 1/8 1/7 vessel Neutrophilsin lumen 1/8 0/8 0/8 0/7 Bacterial embolism, mesenteric blood 0/8 0/80/8 1/7 vessel Right Ureter Dilatation 4/8 2/8 4/8 6/7 Hyperplasia,transitional epithelium 1/8 0/8 0/8 0/7 Vacuolation, transitionalepithelium 1/8 0/8 1/8 0/7 Inflammation, subacute/chronic, peri- 1/8 3/86/8 1/7 ureter mesentery Vasculitis, mesenteric blood vessel 4/8 1/8 2/81/7 Lymph Node, Lumbar Hyperplasia, lymphoid 1/3 0/3 1/2 U Hemorrhage0/3 1/3 0/2 U Histiocytosis, sinus 1/3 3/3 1/2 U Infiltrate, neutrophils2/3 1/3 0/2 U Depletion, lymphocytes 1/3 3/3 1/2 U Lymph Node,Mesenteric Hemorrhage 0/2 1/2 0/2 U Histiocytosis, sinus 0/2 1/2 1/2 UDepletion, lymphocytes 0/2 0/2 1/2 U Congestion 0/2 1/2 0/2 U Adhesionsand Fistulas Ahesion conduit to intestines 8/8 8/8 5/8 5/7 Fistula(macroscopic or clinical notes) 5/8 7/8 6/8 2/7 Fistula/neutrophil tract(microscopic) 3/8 6/8 4/8 1/7 Fistula (macroscopic or microscopic) 5/88/8 6/8 3/7 Adhesion ureter to intestines 1/8 1/8 3/8 2/7 Adhesionconduit to spleen 0/8 0/8 0/8 2/7 Adhesion conduit to seminal vesicle0/8 0/8 1/8 0/7 Adhesion conduit to uterus 0/8 0/8 1/8 0/7 Adhesionureter to uterus 1/8 0/8 1/8 0/7 GROSS LESIONS: Abdominal WallHeterotopic bone, near stoma 4/4 3/5 2/2 1/1 (P = present) Inflammation,chronic-active, 1/4 2/5 0/2 0/1 peritoneum Fibrosis, fascia 0/4 1/5 0/21/1 Gastrointestinal Villous atrophy 2/2 0/1 0/1 U Congestion 0/2 1/10/1 U Mucous hyperplasia, stomach 0/2 0/1 1/1 U Liver Inflammation,chronic-active, U 1/1 U U capsule/peritoneum Lung Inflammation,acute/subacute, 1/1 0/1 3/3 U multifocal Bacterial colonies, multifocal1/1 0/1 1/3 U Vasculitis, necrotizing/thrombi 1/1 0/1 0/3 UInflammation, chronic, pleura 0/1 1/1 0/3 U Lymph Node, InguinalHemorrhage 1/1 0/2 U U Infiltrate, neutrophils 1/1 0/2 U U Hyperplasia,lymphoid 0/1 2/2 U U Depletion, lymphocytes 1/1 0/2 U U Histiocytosis,sinus 0/1 1/2 U U Omentum Inflammation, chronic-active U 3/3 U UPancreas Autolysis (P = present) 1/1 2/4 U U Inflammation,chronic-active, 0/1 2/4 U U peritoneum Seminal Vesicle Inflammation,chronic-active (with 1/2 U U U enlargement) Skin, Ear Vasculitis,necrotizing U 1/1 U U Spleen Inflammation, chronic, peritoneal U U U 1/1surface Uterus Uterus adhered to ureters (P = present) 1/1 U U U

Urothelium regeneration is not dependent on the presence of cells in thetest article and was not expected to occur on the surface of the atriumfacing the conduit lumen; therefore, assessment of urinary tissueregeneration was limited to the cranial end of the conduit (Sections 5,9, 13, and 15).

The placement of the test article and the weight of the overlyingabdominal organs may have contributed to poor drainage and detritusbuild-up in the lumen of all groups; however, there were distinctfindings at necropsy in animals receiving a scaffold-only implant. Thepresence of upper urinary tract lesions was associated with intermittentor complete obstructions resulting in back pressure of the urine—asevidenced by the formation of diverticula in the cranial conduit and theincidence of ureter and kidney damage. No evidence of ureteral orconduit stenosis was observed in any group. However, group 4 animals hadincreased incidence of hydroureter and hydronephrosis relative to Group1, 2, and 3 animals (100% and 69% combined, respectively). The absenceof SMC in the scaffold-only test articles appears to have contributed tothe increase in mortality subsequent to obstruction and incompletedevelopment of urinary tissue, leading to the determination that thescaffold-only test article was unsuitable for further development.

Urinary tissue-like regeneration characterized by mucosa, submucosa andsmooth muscle with a fibrovascular stroma was observed after constructtest article implantation regardless of SMC source (i.e. bladder,adipose or blood). In contrast, a reparative process was observedfollowing implantation of the scaffold-only test article characterizedby an abnormal mucosa supported by fibrovascular stroma with limitedsmooth muscle. The extent of urinary-like tissue regeneration in theconstruct groups was influenced by duration of animal survivalpost-implantation.

FIGS. 60-63 provide photomicrographs of the neo-urinary conduit. FIG. 60shows a photomicrograph (Masson's trichome stain) of cranial portion ofthe NUC from Group 2 female animal 11. Urothelium is present over layersof smooth muscle.

FIG. 61 shows a photomicrograph (Masson's trichome stain) of cranialportion of the NUC from Group 1 female animal 1. Urothelium is presentover layers of smooth muscle.

FIG. 62 shows a photomicrograph (Masson's trichome stain) of the NUCnear the left ureter from Group 2 male animal 15. Urothelium is presentover layers of smooth muscle.

FIG. 63 shows a photomicrograph (Masson's trichome stain) of the NUCwall from Group 1 male animal 5. At this level, the wall of the conduitis comprised of fibrous connective tissue. Detritus covers the luminalsurface. This appearance was most frequently observed in the caudalportion of the NUC.

Conclusions

PCV-2 viral infection and partial to full urinary obstruction of theurinary flow as a result of surgical implantation site and abdominalcontent compressing the lumen contributed to 23/24 unscheduled deaths.Findings related to compression of the lumen and urinary obstructionswere adhesions, fistulas, hydroureter, hydronephrosis, andpyelonephritis. An in-life surgical procedure-related complicationcontributed to 1/24 unscheduled death.

Healing and regeneration was observed in portions of conduits derivedfrom any of the construct test articles while healing and repair wasobserved in the conduits derived from the scaffold-only test article,demonstrating that construct implantation resulted in the formation of aconduit having a urinary-like tissue wall composed of mucosa and smoothmuscle layers.

The difference in healing between the construct test articles(regeneration) and scaffoldonly test article (repair) contributed to ahigher incidence of significant renal findings observed with thescaffold-only test article, leading to a determination that thescaffold-only test article was unsuitable for further development.

There were no differences observed in the regenerative process betweenconstruct test articles, suggesting equivalence between SMC sources inpromoting regeneration.

Example 7 In Vivo Implantation of a Neo-Urinary Conduit Scaffold Seededwith SMCs

A 3 month preclinical study using a porcine model to evaluate thepotential of a synthetic scaffold (PGA) and cell-based constructs tocreate a regenerative Neo-Urinary Conduit (NUC) that would allow urineto flow from the ureters to outside the body without evidence of damageto the upper urinary tract or metabolic abnormalities has been performed(following Example 3 protocols).

The feasibility of using SMC-seeded PLGA-based biodegradable scaffolds,or Neo-Urinary Conduits (NUC), to establish an incontinent urinarydiversion was evaluated. Scaffold-only controls and NUC seeded withautologous SMC isolated from blood, fat, or urinary bladders wereevaluated in a percutaneous diversion porcine model for 3 months. Urineoutflow was maintained by early post-operative management of the conduitlumen and stoma.

The constructs composed of smooth muscle cells (SMC) obtained fromblood, fat, or bladder sources regenerated a patent conduit composed ofa urothelial cell lining and smooth muscle layer that did not result inalterations to the upper urinary tract. No evidence was found forelevated creatinine, metabolic abnormalities or altered hematologicalparameters.

NUC diversions developed into conduits composed of regenerated urinarytissues (urothelial cell lining and vascular smooth muscle wall)regardless of cell source. No significant alterations to the upperurinary tract, creatinine elevations, or hematologic or metabolicabnormalities were observed. In contrast, scaffold-only implantedanimals developed patent urothelial lined conduits composed primarily offibrous connective tissue and limited smooth muscle development. Thisgroup also had a high frequency of hydroureter and hydronephrosis. Inboth groups, early post-operative management of the conduit lumen andstoma were required to maintain patency for the study duration.

FIG. 64 shows the histological characteristics of the regeneratedurological tissue forming the neo-bladder conduit was similar regardlessof SMC origin. Analysis was performed at an interim analysis (48-days)and at end of study (3 mo terminal sacrifice).

These studies demonstrate that a synthetic Neo-Urinary Conduit (NUC)seeded with autologous SMC from various sources (blood, fat or bladder)is capable of establishing a patent incontinent urinary diversion forpost-cystectomy management of urine elimination. NBC-implanted animalsexhibited none of the sequelae commonly associated with GI-derivedurinary diversions or scaffold-only implants. Neo-Bladder Conduit mayrepresent an alternative to GI tissue for post-cystectomy management byincontinent urinary diversion.

According to Pirker et al. (2007) J Urol. April; 177(4): 1546-51, thethree muscle components of the uretero-vesical junction (UVJ) werereadily distinguishable in porcine fetuses at gestational age 60 days.This included the 1) ureteral (U) smooth muscle, 2) detrusor (D) smoothmuscle and 3) the periureteral sheath smooth muscle. The ureteral smoothmuscle was characterized by numerous small diameter muscle fascicles(long arrows), while the detrusor muscle consisted of large diametermuscle bundles running in multiple directions, and the muscle fibers ofthe periureteral sheath distinguishable in all age groups by theirintermediate fiber size, location, and orientation along the intramuralureter (short arrows). U=ureteral lumen; B=bladder lumen, α-SMA stain,reduced from ×40. This is shown in FIG. 65.

As shown in FIG. 66, after 82 days an adipose group animal showedsimilarities between the newly formed uretero-conduit junction (UCJ) andPirker's depiction of the UVJ. Histologically, the UCJ section basicallyshowed similarities to the embryological development of the 60 to 90day-old porcine fetus. Masson's Trichrome-stained histological sectionof UCJ. Original total magnification 10×. U=ureteral lumen; C=conduitlumen, and arrows show a mixture of muscle fibers with ureteral andperiureteral similarities.

FIG. 67 shows the histology of an implanted conduit and indicates thesmooth muscle cell wall, abdominal wall, urothelial cell cover, conduit,and conduit lumen locations.

Example 8 De Novo Functional Neo-Urinary Conduit from Non-Bladder CellSources

We describe here the isolation and characterization of smooth-musclecells from porcine peripheral blood and adipose that are phenotypicallyand functionally indistinguishable from bladder-derived smooth musclecells. We demonstrate that peripheral blood and adipose-derivedsmooth-muscle cells may be used to seed synthetic, biodegradeabletubular scaffold structures and that implantation of these seededscaffolds into a porcine cystectomy model leads to successfulregeneration of a neo-urinary conduit functionally indistinguishablefrom that seeded with bladder-derived smooth muscle cells. The abilityto create urologic structures de novo from scaffolds seeded byperipheral blood- or adipose-derived smooth muscle cells will greatlyfacilitate the translation of urologic tissue engineering technologiesinto clinical practice.

Although smooth muscle cells have also been isolated from other tissuesources such as skeletal muscle and omentum (Wilschut et al. J CellBiochem 105, 1228-1239 (2008); Hernando et al. Eur J Vasc Surg 8,531-536 (1994)), we chose to focus on recovery of smooth muscle cellsfrom peripheral blood and adipose, as these represent the source tissuewith favorable potential clinical utility in terms of the ease of samplecollection. A porcine cystectomy model was selected to evaluate theperformance of peripheral blood- and adipose-derived smooth muscle cellsrelative to bladder-derived smooth muscle cells upon application in acell/scaffold composite (Baldwin et al. J. Endourol. 17, 307-312 (2003);Akbal et al. J Urol. 176, 1706-1711 (2006).

FIG. 68 shows porcine (A) bladder-, (B) adipose-, and (C) peripheralblood-derived smooth muscle cells. Smooth muscle cells from all threetissue types share morphological features characteristic of fullydifferentiated smooth muscle cells, including a flattened,spindle-shaped fibroblastic appearance and whirling, “hill-and-valley”organization.

FIG. 69 shows RT-PCR analysis of smooth muscle cell associated markersfrom porcine bladder-, adipose-, and peripheral blood-derived smoothmuscle cells. Expression of the smooth muscle cell associated markersSMαA, SM22, myocardin, SMMHC and calponin is comparable in all smoothmuscle cell types, regardless of tissue source. Samples are normalizedby mass of RNA and expression of β-actin. Numbers refer to individualswine from which primary cell cultures were derived.

FIG. 70 shows immuno-fluorescence analysis of smooth muscle cellassociated markers from porcine bladder, adipose & peripheralblood-derived smooth muscle cells. Expression of the smooth muscle cellassociated markers SMαA, SM22, SMMHC and calponin is comparable in allsmooth muscle cell types, regardless of tissue source, with theexception of SM22 in adipose-derived SMC Negative controls (IgGisotypes) showed no detectable staining (data not shown).

Direct plating of the peripheral blood-derived mononuclear fraction fromswine resulted in outgrowth of colonies with typical smooth muscle cellmorphology (FIG. 68). All (100%) animals screened (n=24) generatedsmooth muscle cell colonies, with 2.44×10³-2.37×10⁶ smooth muscle cellsrecovered at passage zero from 50 ml of peripheral blood. Recovery ofsmooth muscle cells was unaffected by changes in media formulation, celldensity or surface coatings (data not shown). A similar approach wasused to investigate the potential application of subcutaneous orlipoaspirate-derived adipose as a source of smooth muscle cells. Thestromal-vascular fraction (SVF) of adipose represents a heterogenouspopulation of cells including endothelial cells, smooth muscle cells aswell as progenitor cells with limited mesenchymal potential²². We wereable to generate colonies (expandable into monolayers) of smooth musclecells from porcine adipose with 100% efficiency (n=24), (FIG. 68) with acell recovery rate of 1.37×10⁵-4.36×10⁵ cells/g adipose tissue. Incomparison, smooth muscle cells could be isolated from bladder tissuewith a recovery rate of 1.29×10⁶-9.3×10⁶ cells/g bladder tissue.Expansion of smooth muscle cell colonies from peripheral blood oradipose resulted in the formation of a cell monolayer with a typicalwhirled, “hill-and-valley” organization characteristic of culturedbladder-derived smooth muscle cells (FIG. 68).

Enrichment of smooth muscle cells was facilitated by use of high celldensities and high glucose media, which has been shown to specificallyselect against the growth and expansion of mesenchymal stem cells (Lundet al. 2009 supra). To this end, a systematic comparative analysisdemonstrating the key differences in phenotypic and functionalproperties between adipose-derived smooth muscle cells and mesenchymalstem cells was performed (see Example below).

Analysis of the functional properties of peripheral blood oradipose-derived smooth muscle cells in vitro demonstrates that they areindistinguishable from bladder-derived smooth muscle cells. Increasedexpression of proteins associated with smooth muscle contractility is acharacteristic feature of smooth muscle cell differentiation andmaturation (Jeon et al, 2006 supra; Ross et al., 2006 supra; Sinha etal., 2004 supra). Myocardin is a key transcription factor required forsmooth muscle cell differentiation and acts to mediate the expression ofsmooth muscle markers essential for contractility including SM22,α-smooth muscle actin (SMαA), smooth muscle myosin heavy chain (SMMHC)and calponin (CNN). Expression of the smooth muscle markers SMMHC andCNN is generally regarded as diagnostic of mature smooth muscle cells.(Qiu et al., 2005 supra; Wang et al., 2003 supra; Yoshida et al., 2003supra). As shown in FIG. 69, semi-quantitative RT-PCR analysis of theexpression of these key smooth muscle cell markers demonstrates thatblood and adipose-derived smooth muscle cells are directly comparable tobladder-derived smooth muscle cells.

These results were confirmed by immuno-fluorescence analysis of smoothmuscle cell specific protein expression. αSMA, SM22, CNN and SMMHC wereexpressed in peripheral blood and adipose-derived smooth muscle cellswith a localization pattern identical to that observed inbladder-derived smooth muscle cells (FIG. 70). Localization to stressfibers was observed as is typical for bladder-derived smooth musclecells. Staining of SM22 was observed to be weak in adipose-derivedsmooth muscle cells.

FIG. 71 show contractility of porcine (A) bladder-, (B) adipose-, and(C) peripheral blood-derived smooth muscle cells. Smooth muscle cellsfrom all three tissue sources show Ca²⁺ dependant contractility in acollagen gel matrix. Numbers refer to cell lines derived from individualanimals

The functionality of peripheral blood and adipose-derived smooth musclecells was further evaluated by a three dimensional Ca²⁺-dependantcontractility assay. Smooth muscle cells spontaneously inducecontraction of a collagen matrix in a Ca²⁺-dependant manner whenembedded in a three-dimensional gel (Travis et al. 2001 supra). As shownin FIG. 71, while sample-to-sample variation was observed, peripheralblood and adipose-derived cells contract to a degree comparable tobladder-derived smooth muscle cells and this contractility is inhibitedby EDTA, a known Ca²⁺ chelator.

FIG. 72A-C shows the growth kinetics of porcine (A) bladder-, (B)adipose-, and (C) peripheral blood-derived smooth muscle cells. Thenumbers refer to cell lines derived from individual animals.

Application of adipose and peripheral blood-derived smooth muscle cellsfor urologic regenerative medicine is contingent on being able to secureadequate cell numbers within an acceptable time frame. Towards this end,we have observed that smooth muscle cell colonies (from a 50 ml sampleof porcine peripheral blood or 7-25 g porcine adipose) are identifiablewithin 7 days post seeding, and may be passaged within 14 days. As shownin FIG. 72A-C, with the exception of one bladder smooth muscle samplewhich failed to proliferate for unknown reasons, one million to tens ofmillions of smooth muscle cells were recovered from bladder, peripheralblood or adipose within 2-4 weeks (n=24). Bladder and adipose-derivedsmooth muscle cells were expanded for 2 passages prior to harvesting ofcells for seeding a synthetic, neo-urinary conduit scaffold. Peripheralblood-derived smooth muscle cells were expanded for 3-4 passages togenerate equivalent cell numbers. On average, 30-40×10⁶ smooth musclecells were used to seed a neo-urinary conduit scaffold.

We have previously shown that bladder-derived smooth muscle cells may beused to seed a synthetic, biopolymer scaffold which upon implantationinto an in vivo clinical model of bladder cystectomy resulted in theregeneration of a fully functional de novo bladder augment (Jago Isupra). However, because use of bladder-derived smooth muscle cells maynot be clinically ideal, we proceeded to evaluate the in vivo clinicalefficacy of peripheral blood and adipose-derived smooth muscle cells ina 3 month porcine clinical model of urinary incontinence Baldwin et al.2003 supra; Akbal et al. 2006 supra). Bladder, adipose and peripheralblood-derived smooth muscle cells were used to seed PGA/PLGA-basedscaffolds to create a regenerative, neo-urinary conduit permittingefflux of urine from the ureters directly to the external body surface.We observed that constructs composed of smooth muscle cells obtainedfrom blood or bladder sources regenerated a patent conduit composed ofan urothelial cell lining and smooth muscle layer that did not result inalterations to the upper urinary tract. No evidence was found forelevated creatinine, metabolic abnormalities or altered hematologicalparameters.

FIG. 73 shows the regeneration of neo-urinary conduit in porcinecystectomy model with adipose-, peripheral blood-, and bladder-derivedsmooth muscle cell-seeded synthetic scaffolds. Neo-urinary conduitcomposites seeded with adipose-, peripheral blood-, or bladder-derivedsmooth muscle cells led to regeneration of urinary-like tissue withurothelial and smooth muscle layers. Smooth muscle cell bundles wereobserved in the cranial dorsal/ventral aspect of the neo-urinaryconduit, adjacent to the ureteral/scaffold junction. In all groups,fibroblasts and smooth muscle cells were observed in representativesections obtained from mid-portion of the conduit. The scaffold-onlygroup developed neo-urinary conduits composed principally of fibrousconnective tissue and limited smooth muscle bundles associated withrepair. Masson's trichrome stain was used to identify regenerated smoothmuscle bundles (red) and the presence of early cellular organizationconsisting of fibroblasts and smooth muscle cells in a collagen richmatrix (blue). Nuclei stain dark brown. In all groups, fibroblasts andsmooth muscle cells were observed in representative sections obtainedfrom mid-portion of the conduit. Higher concentration of collagenobserved in the bladder and scaffold groups respectively, characterizedby blue staining from Masson's trichrome stain. Scaffold only groupdeveloped neo-urinary conduits composed principally of fibrousconnective tissue and limited smooth muscle bundles.

As shown in FIG. 73, the histological characteristics of the regeneratedurological tissue forming the neo-urinary conduit was generally similarregardless of the origin of the smooth muscle cell population. Incontrast, scaffold-only implanted animals developed patenturothelial-lined conduits composed primarily of fibrous connectivetissue and limited smooth muscle development. In both groups, earlypost-operative management of the conduit lumen and stoma was required tomaintain patency for the study duration.

This study demonstrates that a synthetic, biodegradable scaffoldcomposite seeded with autologous smooth muscle cells derived frommultiple potential cell sources (blood, fat or bladder) is capable ofbeing used to re-create a patent neo-urinary conduit in a preclinicalporcine model. This ability to create urologic structures de novo fromsynthetic scaffolds seeded by peripheral blood or adipose-derived smoothmuscle cells will greatly facilitate the translation of urologic tissueengineering technologies into clinical practice.

Materials and Methods.

Generation of smooth muscle cells from porcine bladder, adipose andperipheral blood. Smooth muscle cells were isolated from bladder &adipose biopsies as well as peripheral blood draws for use in generationof an autologous Neo-Urinary Conduit construct. A 1 cm² bladder biopsyspecimen, 2 cm² adipose biopsy specimen, and 50 mL of peripheral bloodwas obtained from each of 24 Gottingen swine approximately 8 weeks priorto the planned implantation of the final Neo-Urinary Conduit.

For isolation of bladder-derived smooth muscle cells, the urothelialcell layer was dissected away from the bladder biopsy and the remainingsmooth muscle layer cut into 1 mm² pieces and arranged onto the surfaceof a tissue culture plate. Biopsy pieces were dried in a biosafetycabinet for 10-30 minutes. DMEM-HG (Gibco)+10% FBS was added to thebiopsy samples and the plates incubated in a humidified 37° C. incubatorat 5% CO₂.

Adipose tissue (7-25 g) was washed 3 times with PBS, minced with ascalpel and scissors, transferred into a 50 mL conical tube andincubated at 37° C. for 60 minutes in a solution of 0.3% collagenase(Worthington) and 1% BSA in DMEM-HG. The tubes were either continuallyrocked or periodically shaken to facilitate digestion. Thestromal-vascular fraction was pelleted by centrifugation at 600 g for 10minutes and resuspended in DMEM-HG+10% FBS. The stromal-vascularfraction was then used to seed passage zero.

25 ml of porcine peripheral blood was diluted 1:1 in PBS and layeredwith 25 ml Histopaque-1077 (Sigma) in a 50 mL conical tube. Followingcentrifugation (800 g, 30 min), the mononuclear fraction was collected,washed once with PBS and resuspended in α-MEM/10% FBS (Invitrogen) toseed passage zero.

Assembly of a Neo-Urinary Conduit cell/scaffold composite. Bladder,adipose and peripheral blood-derived smooth muscle cells were expandedseparately for up to 7 weeks to generate the 10⁷ cells required forseeding a NUC scaffold. Bladder and adipose-derived smooth muscle cellswere expanded for 2 passages before harvesting of cells for seeding ofscaffolds to produce the final construct. Peripheral blood-derivedsmooth muscle cell cultures were expanded to P3-4 before harvesting forscaffold seeding. To make the NUC scaffold, PGA felt was cut to size,sutured into the shape of a NUC, and coated with PLGA. This constructwas then sterilized using ethylene oxide. On the day prior to cellseeding, the NUC scaffold was serially pre-wetted by saturation with 60%ethanol/40% D-PBS, 100% D-PBS, D-MEM/10% FBS or α-MEM/10% FBS followedby incubation in D-MEM/10% FBS or α-MEM/10% FBS at room temperatureovernight. The NUC scaffold was then seeded with bladder-, adipose-, orperipheral blood-derived smooth muscle cells and the seeded constructmatured in a humidified 37° C. incubator at 5% CO₂ until implantation inan autologous host pig by day 7.

Isolation of RNA and semi-quantitative RT-PCR analysis. RNA was isolatedfrom porcine bladder, adipose and peripheral-blood derived smooth musclecells using the RNeasy Plus RNA Mini isolation kit (Qiagen). 1 μg of RNAfrom each sample was reverse-transcribed using the Quantitect cDNAsynthesis kit (Invitrogen). The following smooth muscle cell specificprimers were used to set up RT-PCR reactions (5′-3′): β-actin (F: TTCTAC AAT GAG CTG CGT GTG (SEQ ID NO: 1), R: CGT TCA CAC TTC ATG ATG GAG T(SEQ ID NO: 2)), SM22 (transgelin) (F: GAT CCA ACT GGT TTA TGA AGA AAG C(SEQ ID NO: 3), R: TCT AAC TGA TGA TCT GCC GAG GTC (SEQ ID NO: 4)), SMαA(F: CCA GCA GAT GTG GAT CAG CA (SEQ ID NO: 5), R: AAG CAT TTG CGG TGGACA AT (SEQ ID NO: 6)), SMMHC (F: GCT CAG AAA GTT TGC CAC CTC (SEQ IDNO: 7), R: TCC TGC TCC AGG ATG AAC AT (SEQ ID NO: 8)), CNN (calponin)(F: CAT GTC CTC TGC TCA CTT CAA C (SEQ ID NO: 9), R: CCC CTC GAT CCA CTCTCT CA (SEQ ID NO: 10)), MYOCD (F: AAG AGC ACA GGG TCT CCT CA (SEQ IDNO: 11), R: ACT CCG AGT CAT TTG CTG CT (SEQ ID NO: 12)). Cyclingconditions: denature 95° (2 min), denature 95° (45 s), anneal (45 s),extension 72° (45 s), final extension 72° (5 min). 35 cycles (myocardin40 cycles). Annealing temps: β-actin=58°, SM22=56°, SMαA=55°, SMMHC=60°,CNN=51°, MYOCD=52°. PCR reactions were carried out using GoTaq Green PCRmix (Promega) and cycled on an iQcycler (Bio-Rad).

Immuno-fluorescence analysis. The following antibodies were used forimmuno-fluorescence analysis: SMαA (Dako #M0851), CNN (Dako #M3556),SM-MHC (Sigma #M7786), myocardin (Santa Cruz #SC3428), SM22 (Abcam#ab28811-100), anti-msIgG1/Alexafluor 488 (Invitrogen #A21121),anti-msIgG2a/Alexafluor 488 (Invitrogen #A21131), anti-gtIgG/Alexafuor488 (Invitrogen #A11055). All primary antibodies were used at a finalconcentration of 5 □g/ml, except SMMHC which was used at 10 μg/ml.

Contractility assay. Contractility assays were performed as describedpreviously (Travis et al., 2001 supra).

Growth kinetics. Expansion of smooth muscle cells from tissue isolationto seeding of the Neo-Urinary Conduit scaffold was by serial passagingat a confluence ≧70%.

GLP preclinical analysis of de novo neo-urinary conduit formation in aporcine cystectomy model. 32 Gottingen swine with total cystectomy andincontinent ureterostomy (8 animals per data point composed of 4 malesand 4 females each) were used in a GLP preclinical analysis to determinethe safety and functionality of tissue-engineered NUC constructs seededwith autologous smooth muscle cells derived from the bladder, blood oradipose tissue. Of the 32 animals, the first group (4 males, 4 females)was implanted with NUC seeded with bladder-derived smooth muscle cells.A second group was implanted with NUC scaffold seeded withadipose-derived SMCs, a third group was implanted with a NUC scaffoldseeded with blood-derived SMCs, and the 4th group was implanted withunseeded NUC scaffold only. Device effect and performance was monitoredthrough ultrasound imaging, pyelogram, as well as urine and bloodanalysis at different time-points of the study. At the completion of therecovery period (Day 84±5), all animals were euthanized and a necropsyperformed for harvesting the kidneys, conduits, and associated organsand tissues for histological preparation and pathological examination.

Example 9 Assessment of a Neo-Urinary Conduit in Swine

The objective of this study is to determine the safety and functionalityof the use of Tengion's Neo-Urinary Conduit (NUC) Construct seeded withautologous smooth muscle cells derived from the bladder, blood oradipose for conduit implantation and tissue regeneration after surgicalremoval of the bladder (radical cystectomy) and diversion of the uretersto the inflow end of the NUC Construct implant system. Peritonewn willbe used to wrap the whole construct. The draining outflow end of theconstruct will be directed and attached towards the surgically createdstoma in order to pass urine. In this study, the performance of theNeo-Urinary Conduit Construct and the effects on the associated organsand tissues will be evaluated. The endpoint measurements includepyelogram, ultrasound, blood analysis, and histopathology.

Twenty animals (10 F, 10 M) will be enlisted in the study. There is one(I) group with four animals (2 F and 2 M) with autologous bladder SMCs.There are 2 groups with eight animals (4 F, 4 M) each: autologousadipose SMCs and autologous blood SMCs. The first group will beimplanted with a NUC scaffold seeded with bladder-derived SMCs, Thesecond group will be implanted with a NUC scaffold seeded withadipose-derived SMC's, and the third group will be implanted with aNeo-Urinary Conduit scaffold seeded with blood-derived SMCs. Deviceeffect and performance will be monitored through ultrasound imaging,pyelogram, and urine and blood analysis at different timepoints of thestudy. At the completion of the recovery period (Day 84 4-5), theanimals shall be euthanized and a necropsy performed for harvesting thekidneys, conduits, and associated organs and tissues for histologicalpreparation and pathological examination. Four out of twenty animalswill be subjected to two major procedures. The first procedure will be abladder biopsy. At a later date a second surgical procedure will beperformed to implant the neo-urinary Conduit construct seeded withautologous bladder SMCs. The remaining 16 animals will undergo one minorsurgical procedure to collect adipose tissue from the abdomen and venousblood collection. The tissues will be used for harvesting of autologouscells used in constructing the cell seeded Neo-urinary Conduitconstructs. This procedure is required for sufficient autologous tissuesamples needed for the constructs. The two tissues will be collectedfrom the 16 animals to provide the optimal process for group selection.These same animals will be implanted with the autologous SMC seededscaffold derived from the blood or adipose tissues. The utilization ofperitoneum as a vascular source for the conduit implant will also beevaluated.

Test animals: Common Name: Yorkshire Swine; Breed/Class: Sus Scrofa;Number of Animals (by gender): 10 Females & 10 Males; Weight Range: 25±5Kg.

Treatment Groups: The objective of the study is to assess Tengion'sNeo-urinary Conduit Construct in Female and Male Yorkshire pig modelover time for 12 weeks. Table 9.1 shows the five phases of the study.

Summary Phase A Baseline data collection, Tissue harvest for smoothmuscle cells for scaffold Phase B Generation of Test Article (Construct)by Sponsor Phase C Surgical Implantation Procedure of 20 animals Phase DSurvival: Post operative care and monitoring, Observation, DataCollection Phase E Pre-Necropsy follow up & necropsy with tissue harvestand histology

Table 9.2 provides a summary of the study design. The body weight willbe measured before pre-biopsy, pre-surgery, and pre-necropsy. Assessmentof the incision site will be made daily for 14 days or until healed.Maintenance of the stoma button will be daily for the duration of thestudy as needed. Debridement will be performed on an as needed basis peranimal.

TABLE 9.2 No. of Intended Animals Biopsy Surgical ProcedurePostoperative Necropsy Time Group No. Treatment M F Procedure (Day 0)procedures Point 1 Autologous 2 2 Collect from Complete ClinicalApproximately Bladder each animal: cystectomy health 3 months after SMC2.5 cm² tissue followed by assessment, implantation sample transpositionof body weight, (84 ± 5 days the ureters to be stoma button fromimplant) attached to inflow and incision end of NUC site constructmaintenance 2 Autologous 4 4 Collect from Complete blood and adiposeeach animal: cystectomy urine SMC 25-50 g of followed by analysis,adipose tissue transposition of pyelogram sample the ureters to be and 8× 10 ml attached to inflow ultrasound tubes of end of NUC peripheralconstruct blood Ureters are stented with a stent for ~7 days Peritoneumused to wrap and cover whole of construct Draining outflow end of theconstruct is attached through the abdominal wall and exiting to the skinwithout a continent stoma 3 Autologous 4 4 Stoma button Blood SMC placedpermanently to retain patency

Test Devices.

Test Article Group 1: PGA/PLGA Neo-urinary Conduit Construct withautologous bladder-derived SMCs. Description: Scaffold comprising ofsynthetic lactide/coglycolide acid polymers plus autologous bladderderived SMCs.

Test Article Group 2: PGA/PLGA Neo-urinary Conduit Construct withautologous adipose-derived SMCs. Description: Scaffold comprising ofsynthetic lactide/coglycolide acid polymers plus autologous adiposederived SMCs.

Test Article Group 3: PGA/PLGA Neo-urinary Conduit Construct withautologous blood-derived SMCs. Description: Scaffold comprising ofsynthetic lactide/coglycolide acid polymers plus autologous bloodderived SMCs.

Pre-Surgery Fasting

Procedure Description: All animals will be fasted at least 8 hours priorto surgery. Food will be withheld the day prior to each surgery.

Duration/Frequency of the Procedure: Single event the night prior toeach surgery with duration of approximately 8 hours.

Procedure Performance: Food withholding will be performed by DaVINCEsstaff technician in animal pen.

Procedural Records: Data will be recorded on the DaVINCI Animal RoomMaintenance Form, the Twice Daily Assessment Form or Animal ProgressNotes.

Body Weight:

Procedure Description: The weight of the animal will be performed on acalibrated balance.

Duration/Frequency of the Procedure: Pre-Biopsy; Pre-Surgery andPre-Necropsy. Additional body weights may be obtained at the discretionof the Veterinarian.

Procedure Performance: Body weights will be performed by trainedtechnician.

Sedation/Anesthesia:

Procedure Description: Biopsy and Test Device Implantation Procedure:Each animal will be sedated and then anesthetized (in accordance withDaVINCI SOP DAV-SURG-003) prior to surgery preparation. Each animal willbe sedated with Ketamine 20 mg/kg IM, Xylazine 2 mg/kg (FYI) andAtropine (0.04 mg/Kg). Each animal will then be intubated and receiveinhalant isoflurane at 2.5%-4% for induction and 0.5-2.5% formaintenance of anesthesia, delivered through either a volume-regulatedrespirator or rebreathing apparatus. An intravenous sheath will beplaced in a peripheral vessel or right internal jugular vein. LactatedRinger's solution will be administered at 10 mL/Kg/hr for the durationof the surgical procedures (Implantation animals only).

Ultrasound and Other Transient Procedures: Animal will be sedated asdescribed in the paragraph immediately above. Alternatively animals maybe sedated using Telazol (2-5 mg/kg) IM and anesthetized using inhalantisoflurane Of needed) delivered through a cone. At the discretion of theVeterinarian or Surgeon animal may be intubated.

Duration/Frequency of the Procedure: Throughout each day of in-lifeprocedures or surgery

Procedure Performance: Anesthesia will be performed by a DaVINCI surgeonand assisted by trained technicians in DaVINCEs surgery room.

Procedural Records: Data will be recorded on DaVINCI Animal SurgeryForms.

Surgical Preparation:

Procedure Description: For all animals (on biopsy and implantation day),the hair will be clipped from 3 inches above the xiphoid to pubicsymphysis. The animal will then be positioned in dorsal recumbency. Theoperative area(s) will then be cleaned with three alternating scrubs ofpovidone-iodine solution and 70% alcohol; once the alternating scrubsare complete, a final application of povidone-iodine solution will beapplied and allowed to dry. The area(s) will be draped for asepticsurgery.

Duration/Frequency of the Procedure: Prior to Biopsy and prior tosurgery. Surgical preparation will require approximately 30 minutes foreach event.

Urine Sample Collection and Analysis:

Procedure Description: Two urine samples at pre-biopsy and pre-necropsywill be collected by catheterization or test tube caught method.

Urine sample of approximately 3.0 int volume (−1 mL for quantitative and2 int qualitative testing) will be collected in a sterile container forqualitative analysis.

Smaller amounts may be sufficient for analysis; however, if urinecollection is difficult the quantitative sample takes precedent over thequalitative sample. The collected urine will be decanted into 5 inLsterile tubes, refrigerated and shipped 24 hrs of collection.

Qualitative urinalysis: 0.5 mL of urine will be analyzed at DaVINCI onthe day of collection. The qualitative measurement will be taken at timeof collection using Multistixe 10 SG Test Strips.

Quantitative urinalysis: 1 mL or greater will be decanted into 5 mLsterile tubes, refrigerated and shipped in a cooler with an ice packwithin 24 hrs of collection. Parameters of interest for both thequalitative and quantitative measurements include: Qualitativeurinalysis: test strip-Glucose, Bilirubin, Ketones, Specific Gravity,Blood, pH, Protein, Urobilinogen, Nitrites and Leukocytes.

Quantitative urinalysis: qualitative amount of Bacteria, Glucose andtotal protein. Additional parameters may be requested at the discretionof the Facility Veterinarian. Specimens will be analyzed at AniLytics.

Duration/Frequency of the Procedure: Procedure lasts approximately 15minutes per animal. Collection will be done pre-biopsy and prior tonecropsy. Additional urine analyses may be preformed to assess animalhealth at the discretion of the Facility Veterinarian and StudyDirector.

Procedure Performance: Urine collection procedures will be conducted bytrained technicians.

Procedural Records: Collection information and qualitative results willbe transcribed onto a DaVINCI prepared form. Quantitative urine resultswill be reported in hardcopy by analytical laboratory and filed withstudy raw data.

Blood Collection.

Hematology, Coagulation, Serum Chemistry, Blood Gases.

Hematology: Hematology samples will be collected in 2.0 ml EDTA tubes,and stored on wet ice or refrigerated (2-8° C.). Samples will be labeledand analysis will be performed within 24 hrs of collection. Bloodsamples will be evaluated for the parameters specified below: Totalleukocyte count (WBC); Erythrocyte count (RBC); Hemoglobin concentration(HGB); Hematocrit value (HCT) 1; Mean corpuscular volume (MCV); Meancorpuscular hemoglobin (MCH) 1; Mean corpuscular hemoglobinconcentration (MCHC) 1; Platelet count (PLT), wherein 1=Calculatedvalues.

Coagulation: A total of 1.8 mL blood will be collected into 1.8 mLsodium citrate tubes (0.2 mL of 3.8% sodium citrate). Citrated bloodsamples will be kept on ice until ready to be centrifuged at 1,000 to1300×g for 10 to 15 minutes. Before freezing, divide plasma in ½ andfreeze at −70° C. Citrated plasma samples need to be stored at −70° C.Samples will be evaluated for the following parameters: Prothrombin time(PT); Activated partial thromboplastin time (APTT); and Fibrinogen(FIB).

Serum Chemistry: Samples for Serum Chemistry will be collected inapproximately 4.0 ml serum separation tubes. The blood samples will becentrifuged (1300-1600 zg for 10-15 minutes) and the serwn extractedusing sterile technique. Serum will then be frozen at −70° C. Sampleswere evaluated for the following serum chemistry parameters: Glucose(GLU); Urea nitrogen (BUN); Creatinine (CRE); Total protein (TPR);Albumin (ALB); Globulin (GLOB) 1; Albumin/Globulin ratio (A/G) 1;Calcium (CAL); Phosphorus (PHOS); Sodium (NA); Potassium (K); Chloride(CL); Total cholesterol (CHOL); Total bilirubin (TBIL); Triglycerides(TRG); Alanine aminotransferase (ALT); Aspartate aminotransferase (AST);Alkaline phosphatase (ALK); Gamma glutamyltransferase (GGT); where1=Calculated values.

Blood Gases: Blood Gas/Spun Hematocrit/Total Protein Monitoring:Arterial blood samples (−1.0 mL) will be collected and analyzed using acalibrated i-STAT analyzer and the appropriate cartridge. Samples willbe evaluated for the following blood gas parameters: Sodium (Na)(mmol/L) PCO2 (mm Hg); Potassium (K) (mmol/L) PO2 (mm Hg); IonizedCalcium (iCa) (mmol/L) TCO2 (mmol/L); Glucose (Glu) (mg/dL) HCO3(mmol/L); Hematocrit (Hct) (%) BEecf (mmol/L); pH; and SO2(%).Duration/Frequency of the Procedure. Collection of blood samples foreCBC, Clinical Chemistry and Coagulation will be conducted pre-biopsy,weeks 4 & 8 and pre-necropsy. Blood gases will be collected pre-biopsy.Each blood collection procedure lasts approximately 15 minutes peranimal.

Vital Signs Monitoring Procedure:

Procedure Description: The following vitals will be monitored duringimplant surgery: animal will be monitored approximately every 20 minutefor oxygen rate, Sa02, Pulse rate, Respiration, and body temperature.

Duration/Frequency of the Procedure: At approximately 20 minuteintervals throughout the surgical procedure.

Procedure Performance: Vital signs monitoring procedures will beconducted by trained technicians in DaV1NCI's surgery room.

Blood Collection, Adipose Tissue Collection and Bladder TissueCollection for Cell Culture:

Procedure Description: Blood Collection (Group 2 & 3): Approximately8×10 ml aliquots of venous blood collected from sixteen (16) animals (8M, 8 F) in heparinized vacutainers. The blood will be packaged in thesponsor supplied shipper and shipped overnight to the Sponsor forprocessing. Adipose Tissue Collection (Group 2 & 3): To access theAdipose Tissue and the Bladder a midline incision will be made in theabdomen beginning immediately caudal to the umbilicus. From the same 16animals that blood was collected, subcutaneous adipose tissue will beaseptically collected from the abdomen, corresponding to approximately25-50 grams of tissue. The biopsy tissue will be immediately preservedaseptically in sponsor supplied jars containing tissue culture media andthen packaged according to section 14.8 and shipped overnight to theSponsor for processing.

Bladder Tissue Collection (Group 1): In the remaining four (4) animals,the urinary bladder will be exposed and emptied of urine. One apicaldome piece (˜2.5×2.5 cm) of urinary bladder tissue will be excised fromthe bladder. The urinary bladder tissue will be immediately preservedaseptically in the tissue culture media supplied by the Sponsor and thenpackaged according to section 14.8. The defect in the bladder will thenbe closed utilizing an appropriate technique, using absorbable suturematerial.

The abdominal incision will be closed in layers with absorbable suturematerial of an appropriate size. The skin will be closed in asubcuticular fashion, again using an appropriate size of absorbablesuture material.

Duration/Frequency of the Procedure: Once per source animal lastingapproximately 1 hour per animal.

Procedure Performance: Biopsy collection and preservation will beperformed. The blood may be collected either in animal cage area orsurgical suite as aseptic as possible.

Ureteral Transposition Through Conduit with Cystectomy:

Procedure Description Summary: The ureteral transposition procedure willbe performed via laparotomy. A midline incision will be made in theabdomen beginning 5 cm cranial to the umbilicus extending approximately15 cm caudal. The peritoneum will be identified, carefully separatedfrom the abdominal space until the tissue is long enough to cover theNeo-urinary Conduit Construct and form a conduit that can exit throughthe body wall. The peritoneum will be measured and cut in order to wrapthe construct and form a conduit that will extend out of the body wall.The peritoneum will be sutured around the construct with 3-0 Vicryl.Care will be taken to ensure the tissue remains intact and vascularized.The urinary bladder will then be exposed and emptied of urine takingcare to avoid urine from entering into the abdominal cavity. Thearteries and veins supplying the bladder will be identified and ligated.The ureters will be identified, two 7Fr 14 cm non-absorbable ureteralstents (DaVinci made) will be inserted in ascending fashion and theureters will be carefully transected from the bladder. The urethra willbe over sewn as it is transected. The bladder will then be removed. Theleft ureter will be carefully freed from the surrounding retroperitonealfascia extending cranially until there is enough mobility to reach theright side. The right ureter will be dissected free to reach the end ofthe construct. The ureters will be sutured on to the construct with 3-0Vicryl in a simple continuous pattern. A stoma will be created on theventral abdominal wall lateral to the mammary glands. The peritonealconduit will be exteriorized and sutured to the skin. Surgical adhesivewill be placed along the suture line and where the peritoneum exits thebody wall. The suture strands that are connected to the stents will beexteriorized through the stoma for future removal, and a DaVINCI madestoma button/catheter of appropriate length will be inserted into thestoma allowing adequate drainage for the duration of the survivalperiod. Once secured, the abdominal incision will be closed withnonabsorbable Prolene suture. The skin will be closed in a routinefashion. The animal will then be recovered in animal's cage. NOTE: Theperitoneum must be handled and manipulated with great care to preventstaunching blood flow through the vasculature. The ureteralnon-degradable stents will be left in place for approximately 7 daysunless diagnostic evaluations reveal a need to remove them prematurely(e.g., renal obstruction).

Duration/Frequency of the Procedure: Once per animal lastingapproximately 2-4 hours per animal on Day 0.

Stoma Button Care and Maintenance & Incision Site Assessment

Procedure Description:

Stoma Button/Catheter (Foley or Equivalent): After the definitivesurgery, the stoma catheter (DaVINCI generated stoma button orequivalent: 3-10 cm based on needed at various timepoints) will bereinserted and secured to the animal with sutures. Stoma button will bekept in place for the duration of the study. The catheter will beflushed with sterile saline when it is not dripping to assure patency.Note: Between days 7 and 21, scaffold material undergoes degradation andparticulates (protein-associated) start to be shed in the urine. Thismay cause obstruction of the stoma button and retention of urine volumeabove or beyond the construct's capacity. Therefore, debridement of thestoma and or neo-conduit will be conducted as necessary.

Incision Site Assessment: The incision site will be evaluated daily forthe initial 14 days or until healed. The stoma area and surroundingtissue will be cleaned twice a day. Stoma will be observed for urinedrainage, incision site will be evaluated for dehiscence, abnormaldischarge, odor, irritation or any abnormalities.

Stagnant stoma tissue debridement procedure: Animals with stagnanttissue within the stoma/conduit will undergo a debridement procedure.Animal will be sedated according to protocol. A small incision may bemade on the stoma to facilitate insertion of forceps for debridement.The stagnant issue will be visually identified and grasped with forcepsand gently tugged. Once all stagnant tissue is removed, thestoma/conduit may be flushed with saline solution. The incision will beclosed with a suture(s) and the stoma button will be reinserted andsecured to the animal with sutures. Animal will be recovered inindividual cage.

Duration/Frequency of the Procedure: Following surgical procedure asfollows: Stoma Button: Daily observations followed by maintenance asneeded when catheter is observed not dripping. Time required forapproximately 15 minutes. Incision Site Assessment: Daily for theinitial 14 days or until healed and/or at the discretion of the FacilityVeterinarian. Time required for approximately 15 minutes. 14.11.3Procedure Performance: Catheter care and incision site assessment willbe preformed by either the Facility Veterinarian or trained technicians.

Clinical Observations:

Procedure Description: Recovery: Immediately following completion ofeach surgery, animal will be allowed to recover from anesthesia andtransferred to the home cage. Clinical Observations Post-Implantationfor a 4 Week Period: Postimplantation, individual animal evaluations offood intake and fecaUurine output will be conducted daily for 5 days perweek (i.e., Monday Friday) post implantation for 4 weeks. Observationperiod may be extended at the discretion of the Facility Veterinarianand/or the Study Director. Survival: Post-implantation/reimplant surgeryrecovery animals will be survived for a period of 84±5 days.

Duration/Frequency of the Procedure: Recovery: will be performed at theend of any surgical procedure, approximately 1 hour. Clinicalobservations 5 days Post-implantation/reimplant surgery: Clinicalobservations will be conducted daily for 5 days per week.

Daily Animal Health Assessment: will be performed twice per day,approximately 8 hours apart for approximately 10 minutes for theduration of the study from quarantine to necropsy.

Pyelogram and Ultrasound:

Procedure Description:

Pyelogram will be performed through a peripheral vein, or alternatelyvia femoral artery catheterization under fluoroscopy injection ofcontrast directly into the renal artery. Ultrasound will be performedunder general anesthesia on kidneys and Neo-urinary Conduit.

Duration/Frequency of the Procedure:

Pyelogram: Pre-necropsy. Procedures last approximately 30 minutes peranimal. Ultrasound: Pre-biopsy (kidneys), week 4, 8 and pre-necropsy(kidneys and Neo-urinary Conduit). At the discretion of the StudyDirector and the Attending Veterinarian, additional ultrasound imagingmay be obtained at other timepoints as needed to assess the animal'sclinical health.

Animal Sacrifice and Necropsy.

Procedure Description:

Unscheduled Necropsy—Any animal found dead, moribund or undergoing anunscheduled euthanasia will be subjected to a limited necropsy. Theexploratory necropsy will attempt to determine any potential cause(s) ofdeath or issues leading to euthanasia. Tissue collection will be limitedto the urogenital system. Microscopic analysis of the tissue will bedecided through a joint discussion between Study Director and SponsorRepresentative. Unscheduled and Scheduled Euthanasia—All animals will beinjected with sodium pentobarbital (150 mg/kg, IV) to cause euthanasiain accordance with accepted American Veterinary Medical Association(AVMA) guidelines. Scheduled euthanasia is to be at week 12 (day 84±5)postimplantation.

Physical Examination—All animals will be evaluated by the FacilityVeterinarian prior to euthanasia. The examination will include recordingthe general condition of the animal: rectal body temperature,respiratory rate, heart rate, and capillary refill time. Necropsy—Allanimals will be subjected to necropsy. There will be a specific focus onthe kidneys, conduit, ureters, uretero-vesical junctions, midconduit,conduit-skin junction, and lymph nodes (lumbar and mesenteric). Grossevaluation will be performed on the kidneys, ureters, urethra (ifpresent), conduit, stoma, thoracic, abdominal & pelvic cavities andtheir organs and tissues. If any gross lesions, adhesions and/or organchanges (including reproductive) are observed, they will be evaluated,photographed and collected for histopathological assessment. Majororgans of the abdominal cavity will be collected and saved for possiblefuture microscopic analysis. The major organs are the liver, spleen,pancreas, large intestine (cecum, colon, and rectum), small intestine(duodenum, jejunum and ileum) and the stomach (cardia, fundic andpyloric). The complete neo-urinary conduit area will be visualized andphotographed in situ. Additional photographs and/or gross lesion may betaken at the discretion of the prosector. Fixation of conduit will bedone with fonnalin by infusion of fonnalin into the stoma and inflatingthe conduit and ureters. This will be done with Foley (or equivalent)catheter while the stoma is tied off to hold pressure. 14.15.2Duration/Frequency of the Procedure: Single event duration ofapproximately ½ hr per animal on Day 84 (±5 days)

Histology/Pathology Laboratory

Procedure Description: The fixed urinary organs (i.e., implantedneoconduit, kidneys, and associated tissues) will be collected, trimmed,examined, embedded in paraffin, and sectioned. Slides will be stainedwith hematoxylin and eosin (H & E) and Masson's Trichrome (elastin).

Example 10 Formation of a Conduit Having an Epithelialized Mucosa

Following the Example 9 protocols above, animals were implanted with aNUC scaffold seeded with adipose-, peripheral blood-, or bladder-derivedsmooth muscle cells.

Post-operative clinical observations were similar across all threetreatment groups. All animals in all groups had urine flowing from thestoma immediately following post-operative procedures. All animals inall groups were clinically normal by one month. All animals in allgroups required stoma button maintenance and debridement to maintainurine flow from stoma. Serum markers of renal function (blood ureanitrogen [BUN] and serum creatinine) were similar at baseline across alltreatment groups. At week 4, the values increased for all groups (datanot shown). Hematology indicators of inflammation and/or safety concerns(total white blood cells and fibrinogen) were similar at baseline acrossall treatment groups. At week 4, the values increased for all animals(data not shown).

Following necropsy, the fixed urinary organ (i.e., implanted neoconduit,kidneys, and associated tissues) was collected, trimmed, examined,embedded in paraffin, and sectioned. Slides were stained withhematoxylin and eosin (H & E) and Masson's Trichrome (elastin). Theconduits formed in the animals were examined and found to becharacterized by an epithelialized mucosa at the stomal end. As shown inFIG. 74, the epithelialized mucosa is located at the stomal end(Neo-urinary conduit seeded with adipose-derived SMC). AE-1/AE-3 wasused to detect cytokeratin markers as an indicator of epithelium.

Example 11 Neo-Bladder Augmentation Constructs after Trigone-SparingCystectomy

The neo-bladder constructs of this study will be formed from abiodegradable scaffold on which a subject's own organ specific cells aregrown. The goal is to construct a new urinary bladder for the subjectthat can be implanted in place of the diseased one, alleviating the needfor creation of a “cloaca”. The objective of this study is to determinethe equivalency of neo-bladder constructs for urinary bladderaugmentation.

Experimental design. Each of the 6 groups of canines will include 3females and 3 males in which different densities of urothelial cells(UCs) and smooth muscle cells (SMCs) will be tested. Bladder augmentscaffolds will be seeded with cells as follows.

Group Description Cell source 1 10 × 10⁶ UC and 10 × 10⁶ SMC Bladder 20.1 × 10⁶ UC and 20 × 10⁶ SMC Bladder 3 0.01 × 10⁶ UC and 20 × 10⁶ SMCBladder 4 0 UC and 20 × 10⁶ SMC Bladder 5 0 UC and 20 × 10⁶ SMC Blood 60 UC and 20 × 10⁶ SMC Adipose tissue

Tissue biopsy procedures. Groups 1-4 use SMCs and UCs derived from thebladder biopsy. Prior to the date or while the animal is anesthetizedfor the bladder biopsy procedure, each animal will be prepared foraseptic blood collection from a peripheral vessel and approximately 60mL of venous blood will be aseptically collected (six 10 mL blood tubeswith sodium heparin). Once the midline incision for urinary bladderbiopsy is made in the abdomen, adipose tissue will be exposed andexcised. The amount of adipose tissue collected aseptically from theabdomen will correspond to approximately 20-50 grams of tissue. If it isdetermined that there is not a sufficient amount of tissue collected, anincision will be made over the inguinal fat pad and subcutaneous tissuesdissected to obtain adipose tissue. The urinary bladder will be exposed,and the bladder will be emptied of urine, should any be present. One,approximately 2 cm×1 cm piece of urinary bladder will then be excisedfrom the apex of the bladder.

Surgical procedure (day 1)—a trigone-sparing cystectomy is performedfollowed by implantation of a neo-bladder construct. Postoperativeprocedure—cycling, compliance measurement, fluoroscopic examination,general health assessment, and clinical treatment are performed asnecessary. Necropsy time point—approximately 6 months after implantation(182±2 days from implantation).

Animals. The species Canis familiaris (strain—mongrel dog) is used. 12males (plus one alternate) are used. 12 females (plus one alternative),which are nulliparous and nonpregnant are used. Animals are young adultsat the biopsy stage and weigh between 15-25 kg.

Preoperative Procedures.

Analgesia and Anesthesia. Before induction of anesthesia the animal willbe premedicated with atropine SO₄. (0.04 mg/kg, subcutaneously [SC]),buprenorphine (0.01 mg/kg, SC), and meloxicam (0.2 mg/kg, SC). Anintravenous catheter will be placed in a peripheral vessel and at least10 minutes after administering atropine, the animals will beanesthetized with Propofol® (4-10 mg/kg, intravenously [IV]). The animalwill then be intubated and maintained in anesthesia with isofluraneinhalant anesthetic, delivered to effect through a volume-regulatedventilator. Lactated Ringer's solution will be administered during theprocedure at a rate of approximately 5-20 mL/kg/hr. Each animal willreceive a bupivacaine line block at the site of the abdominal incision(≦2 mg/kg, SC).

Antibiotic Therapy. To help prevent infection, the animals will be givencefotaxime (50 mg/kg, IV) immediately before the biopsy procedure.Additional doses may be administered as necessary. At the time of thedefinitive operation (Day 1), the animals will be given cefotaxime (50mg/kg, IV) immediately before surgery and at its completion.

Animal Preparation. Lacrilube® (or other suitable ocular lubricant) willbe applied to both eyes. Animals will be kept warm throughout thepreparation and the surgical procedures. Hair will be clipped from theentire ventral abdomen. The surgical site will be prepared for asepticsurgery by first wiping the area with povidone-iodine scrub solution toremove all detritus, followed by wiping the area with sponges that havebeen soaked in 70% isopropyl alcohol. The area will then be allowed todry. The animal will be taken to the operating room and placed in dorsalrecumbency. A temperature probe will be inserted into the esophagus inorder to monitor core temperature. At the time of biopsy surgery, theurinary bladder will be catheterized with a dual lumen catheter (orequivalent) to obtain a baseline compliance measurement. The catheterwill be removed from the animals after the measurement. At the time ofdefinitive surgery, the urinary bladder will be catheterized with aFoley catheter. Before both procedures, the surgical sites will again bewashed thoroughly with povidone-iodine scrub solution, wiped withsponges that have been soaked in 70% isopropyl alcohol, and then allowedto dry. DuraPrep™ (or similar) solution will be applied to the area andalso allowed to dry. The area will then be appropriately draped forstrict aseptic surgery.

Surgical Procedures

Urinary Bladder Biopsy. A midline incision will be made in the abdomen,beginning immediately caudal to the umbilicus. The urinary bladder willbe exposed, and the bladder will be emptied of urine, should any bepresent. One, approximately 2 cm×1 cm piece of urinary bladder will thenbe excised from the apex of the bladder. The urinary bladder tissue willbe preserved in tissue culture media (DMEM or equivalent). The defect inthe bladder will be closed in at least 2 layers, using absorbable suturematerial (PDS or equivalent). Alternatively, the bladder will be closedwith surgical staples, oversewn with suture (PDS or equivalent). Theabdominal incision will be closed in layers with absorbable suturematerial of an appropriate size. The skin will be closed in asubcuticular fashion, again using an appropriate size of absorbablesuture material. Alternatively, the skin may be closed with staples.

Neo-bladder augment construct implantation. Neo-bladder augmentscaffolds are seeded with UCs and/or SMCs as described above to formneo-bladder augment constructs for implantation. A midline incision willbe made in the abdomen, beginning immediately caudal to the umbilicus. Aself-retaining abdominal retractor may be placed to open the incision.The urinary bladder will be exposed, and the bladder will be emptied ofurine, should any be present. The area of the trigone will beidentified, and the urinary bladder will be resected in toto but leavingthe trigone and ureteral valves intact. A catheter will then be advancedinto what will be the construct's lumen by passing it through asubmucosal/subserosal tunnel in the trigone region. This catheter willthen be secured to the bladder serosa with the appropriate suturematerial, and will be brought to the outside of the animal by tunnelingit through the abdominal musculature, subcutaneous tissues and skin,exiting near the umbilicus.

The construct will be anastomosed to the normal urinary bladder tissueusing a series of suture patterns to be determined at the actual time ofsurgery with polyglactin 910 suture material. The lateral margins (bothright and left) of the anastomotic site will be marked with anonabsorbable suture to aid in identifying the anastomotic line at thetime of necropsy. The omentum will be pulled over the bladder constructand secured with surgical adhesive.

The abdominal incision will be closed in layers with absorbable suturematerial of an appropriate size. The skin will be closed in asubcuticular fashion, again using an appropriate size of absorbablesuture material. Alternatively, the skin may be closed with staples. TheFoley catheter will be left in place to facilitate postoperative urinecollection after the urinary bladder augmentation procedure with thetest device.

Incision Site Observations. The surgical incision(s) will be observedand assessed at least once daily for at least 14 days (or until healed)for signs of infection, inflammation, and general integrity aftersurgery. Skin staples (if used) may be removed between 7 and 21 daysafter surgery. Appropriate therapy will be initiated as necessary.

Postoperative Urine Collection. After the definitive surgery, urine willbe collected from the catheters until they are removed. All catheterswill be connected to negative pressure reservoirs to collect urine fromthe animals. The reservoirs will be emptied as needed and the volumewill be recorded. The reservoirs will be appropriately attached to theanimal using a canine jacket, or equivalent, to prevent its disruption.The dogs may wear an Elizabethan or similar collar until adequatelyhealed and continence has been achieved.

In the females, the perineal area, vulva, and vagina will be cleanedtwice a day, and the vagina will be aseptically infused with antibioticcream twice a day. The preputial area in the males will be similarlycleaned. In addition, the exit sites for the remaining catheters will beexamined and cleaned at least twice daily. Catheters will be flushed asneeded. If required, due to the suspected presence of urine in theperitoneal cavity, peritoneal lavage may be performed.

Appropriate measures to prevent disruption of the catheters and urineleakage will be performed during this period. The indwelling urethralcatheter (ie, Foley) may be removed within approximately 7 days ofplacement. The suprapubic/subserosal/percutaneous catheter will beremoved approximately 14-21 days after surgery. Once the Foley andsuprapubic catheters have been removed, urine may be collected bycatheterization or by a “pan caught” method.

Compliance Measurement. Before the biopsy procedure, monthly beginningapproximately 30±3 days after implantation, and on the day of necropsy,a compliance measurement will be obtained. The urinary bladder will becatheterized with a dual-lumen catheter. All residual urine will beremoved and the catheter size and placement length will be recorded toassure consistency for follow-up procedures. One lumen will be connectedto the direct pressure cable and the other lumen will be used to infusesterile saline (warmed by incubator) at a rate of 10-25 mL/min. Thestarting pressure of 0-10 mmHg will be achieved and recorded along withthe start time. Time, volume delivered, and the pressure obtained willbe recorded at the time leakage is observed around the catheter (akaleak point). The total infused amount of saline will then be aspiratedto empty the bladder (either completely or partially depending on ifcycling or fluoroscopic imaging follows) and the volume recovered willbe recorded. Animals will be tranquilized as outlined in Section 13.7.If a leak pressure of zero is obtained, the measurements will berepeated at least once, but not more than 3 times.

Cycling. Cycling will be performed every 2 weeks (14±2 day intervals)starting approximately 1 month after implantation and continuing untilapproximately Day 90. Cycling will be completed after compliancemeasurement and before fluoroscopic imaging. Cycling will be performedby re-inflating the bladder with sterile saline (warmed by incubator)after the completion of compliance measurement at a rate of 10-25mL/min. The cycling will be repeated at least 5-10 times. The startingpressure of 0-10 mmHg will be achieved and recorded along with the starttime. Time, volume of isotonic solution delivered, and the pressureobtained will be recorded for each cycle at the time leakage is observedaround the catheter (aka leak point), or when the volume delivered isequal to that of the compliance measurement just performed, whichevercomes first.

Fluoroscopic Imaging. Fluoroscopic imaging will be performed oncemonthly beginning approximately 30±3 days after implantation and on theday of necropsy. The fluoroscopic imaging will be conducted by infusingcontrast media into the bladder and recording it. Fluoroscopic imagingwill be performed after the completion of the compliance measurement orcycling (as appropriate to time point). Approximately half of the totalinfused saline will be aspirated and replaced with a 50/50 mixture ofsterile saline (warmed by incubator) and a contrast media to inflate tothe most recent leak volume of the bladder. Fluoroscopic imaging will beperformed throughout the infusion of the 50/50 mixture. The volume usedfor the cystogram will be that volume at which the leak-point pressurewas obtained.

In-Life Observations and Measurements.

Moribundity/Mortality Check. Moribundity/mortality checks will beperformed twice daily (AM and PM). All animals will be checked forgeneral health, mortality, and moribundity.

Clinical Observations. After the biopsy procedure, clinical observationswill be performed at least once weekly. After the implantation, clinicalobservations will be performed at least twice daily (at least 6 hoursapart) for the first 2 weeks and then daily until Day 30. Clinicalobservations will be continued at least once weekly (7±1 day)thereafter. All animals will be observed; observations will be recorded.

Body Weights. Body weights will be recorded before animal assignment,within 5 days before the biopsy procedure, within 5 days beforeimplantation, weekly thereafter (7±1 day) for the first 3 months (ie,until Day 90), and then monthly (intervals of 30±2 days) until necropsy.

Physical Examinations. A physical examination, including a record ofgeneral condition, rectal body temperature, respiratory rate, heartrate, and capillary refill time, will be performed for each animalbefore entry into the study and before necropsy.

Concurrent Therapy. In accordance with accepted veterinary practices,the animals may be administered concurrent therapy (such as antibioticsor fluid therapy) as required to maintain general good health of theanimals. Concurrent therapy will be administered as necessary.

Sample Collection.

Blood. Blood will be collected from a peripheral vessel. Blood volumesrepresent whole blood and are approximate amounts. The following BloodSample Collection Schedule will be followed.

Serum Time Point Hematology Chemistry Coagulation Before biopsy and X XX implantation (before Day 1) Once weekly for first month X X X afterimplantation (7 ± 1 day intervals), every 2 weeks thereafter (14 ± 2 dayintervals), and on the day of necropsy Volume of Whole Blood 0.75 mL 1.1mL 1.3 mL Anticoagulant EDTA None Sodium Citrate

Urine. Urine samples will be collected via the Foley catheter (while itis indwelling), by catheterization after the Foley catheter has beenremoved, or by “pan caught” method. After collection, samples will betransferred to the appropriate laboratory for processing and analysis.The following Urine Sample Collection Schedule will be followed.

Time Point Urine Before biopsy and before implantation (Day 1) X Onceweekly for first month after implantation X (7 ± 1 day intervals), every2 weeks thereafter (14 ± 2 day intervals), and on the day of necropsyVolume of Urine 3.0 mL Anticoagulant None

Clinical Pathology

Hematology. Blood samples will be evaluated for the followingparameters: Red blood cell count; Hemoglobin concentration; Hematocrit;Mean corpuscular volume; Mean corpuscular hemoglobin concentration; Meancorpuscular hemoglobin; Reticulocyte count; Red blood cell morphology;Platelet count; Platelet morphology; White blood cell count; Neutrophilcount; Lymphocyte count; Monocyte count; Eosinophil count; Basophilcount; Other cells (as appropriate)

Coagulation. The blood samples will be centrifuged, the plasma will beextracted, and the plasma samples will be evaluated for the followingparameters: Coagulation Parameters; Activated partial thromboplastintime; Prothrombin time; Fibrinogen.

Serum Chemistry. The blood samples will be centrifuged, the serum willbe extracted, and the serum samples will be evaluated for the followingparameters: Serum Chemistry Parameters; Alanine aminotransferase;Aspartate aminotransferase; Alkaline phosphatase;Gamma-glutamyltransferase; Total bilirubin; Blood urea nitrogen (BUN);Creatinine; Calcium; Phosphorus; Total protein; Albumin; Globulin;Albumin/globulin ratio; Glucose; Cholesterol; Triglycerides; Sodium;Potassium; Chloride.

Urinalysis. Urine samples will be evaluated for the followingparameters: Volume; Color; Clarity; Specific gravity; Microscopicevaluation of urine sediment; Urine test strip analysis, including: pH,Protein, Glucose, Bilirubin, Ketones, Blood, Urobilinogen, Nitrites,Leukocytes.

Euthanasia. On the day of euthanasia, the animals will initially betranquilized and the compliance measurement and fluoroscopic imagingwill be performed. Euthanasia (deep anesthesia of sodium pentobarbital,35-60 mg/kg, intravenously, to effect, followed by exsanguination) willthen be performed.

Gross Necropsy. A complete gross necropsy will be conducted on allanimals. The necropsy will include examination of the carcass andmusculoskeletal system, all external surfaces and orifices, cranialcavity and external surface of the brain, and all thoracic, abdominal,and pelvic cavities with their associated organs and tissues. There willbe specific focus on the urinary bladder.

Tissue Collection and Preservation. The abdominal cavity will be openedand the augmented urinary bladder will be visualized and photographed insitu. The complete bladder will then be removed (trigone, anastomoticsite, and neo-bladder), the ureters ligated, and the urethraappropriately catheterized to allow it to be fixed under pressure,similar to those generated during compliance measurements, with theappropriate fixative (ie, 10% neutral buffered formalin [NBF]) for themethods of evaluation (histopathology). After fixation in 10% NBF for21-48 hours, the tissues will be transferred to 70% ethanol. Inaddition, organs (or samples of organs) and tissues listed below will beexamined in situ, dissected free, and fixed in 10% NBF or other suitablefixative.

The following tissues will be collected: Adrenal gland (paired); Animalidentification (collect at necropsy to retain identification); Aorta;Bone marrow; sternum; Brain (cerebrum, cerebellum, brain stem); Cervix;Epididymis (paired); Esophagus; Eye (paired) (Fixed in Davidson'sSolution); Gallbladder; Heart; Intestine, cecum; Intestine, colon;Intestine, duodenum; Intestine, ileum (with Peyer's patch); Intestine,jejunum; Intestine, rectum; Kidney (paired); Lacrimal gland (paired);Liver; Lung; Lymph node, mandibular; Lymph node, mesenteric; Lymph node,iliac; Mammary gland; Nerve, optic (paired) (Fixed in Davidson'sSolution); Nerve, sciatic; Ovary (paired); Pancreas; Parathyroid glandPituitary gland; Prostate gland; Salivary gland, mandibular (paired);Skeletal muscle; Skin; Spinal cord (cervical, thoracic, lumbar); Spleen;Stomach (cardiac, fundic, pyloric); Testis (paired) (Fixed in ModifiedDavidson's Solution); Thymus; Thyroid gland (paired); Tongue; Trachea;Uterus; Vagina; Gross lesions/masses (Fixed in Modified Davidson'sSolution).

Histology. The fixed urinary neo-bladder will be cut in half creating adorsal half and a ventral half (the ureters enter the bladder on thedorsal surface). The bladder will again be cut in half along acranial/caudal line creating 4 quadrants of bladder tissue. From eachquadrant of tissue, 3 samples of up to ˜0.5-cm wide will be collected(full thickness). As an identification aid, samples will decrease inlength from Sample 1 (longest) to Sample 3 (shortest). In addition, toaid in orientation of tissue, a nick incision will be made in theserosal/adventitial portion of the caudal end of each sample.

When the surgical interface is apparent, two samples (i.e., Samples 1and 2) from each quadrant will be collected from areas that span thesurgical interface of the native bladder and neo-bladder. These sampleswill come from the area just above where the ureter inserts into thebladder (outside not inside). A third sample (ie, Sample 3) from eachquadrant will be collected from the area cranial to the surgicalinterface representing the urinary neo-bladder samples and may not spanthe surgical interface.

When surgical interface is not apparent, three samples will be collectedin a linear fashion from caudal (i.e, trigone) to cranial (i.e., apex).

A total of 12 samples will be collected from each augmented urinarybladder. Tissue samples will be trimmed, embedded in paraffin, andsectioned for histology evaluation. Regardless of sectioning scheme, the3 samples from each quadrant will be embedded in 1 block so that thereshould be 4 blocks per bladder. The tissue samples will be embedded insuch a way that when they are sectioned, the sections go through alllayers of the tissue rather than en face (such as the epithelium only).

Slides for histopathology will be stained with hematoxylin and eosin (H& E) and Masson's Trichrome (elastin).

In addition, the ureters, urethra, kidneys, representative trigonesamples, and local lymph nodes will be trimmed, embedded in paraffin,and sectioned. Slides will be stained with H & E.

Bladder Wall Thickness. Quantitative measurements of the wall thicknessof each section will be performed with intra- and inter-groupcomparisons assessed. Blinded measurements of the primary constituentparts of the bladder wall, the combined urothelium/lamina propria(Uro/LP), and the tunica muscularis (TM), will be performed manuallyusing an ocular micrometer at 4× magnification and denoted as reticules.The reticules will then be converted to millimeters and the averagethicknesses of the Uro/LP, TM, and total wall will be calculated. Thosesections which had been interpreted as approximating anatomic normalcy(ie, ‘anatomic’) during the histologic grading phase will be assessedfor comparative analysis. These measurements will then be used tocompare the relative thickness of the 3 noted regions of the bladder(ie, the trigone, mid, and apical bladder regions) as well as assess theoverall consistency of the total wall thickness and the ratio andpercentage of the constituent parts thereof. All values will bepresented in the form of averages.

Statistical analysis. Data will be presented as individual values byanimal and summary values with calculated means and standard deviations.Statistical analysis will be performed on body weights, hematology,coagulation, and serum chemistry. To determine the appropriatestatistical test, each data set will be subjected to a statisticaldecision tree using the SAS® System. A minimum of 3 animals per sex pergroup per interval will be required for statistical analysis. The datawill initially be tested for normality using the Shapiro-Wilk testfollowed by the Levene's test for homogeneity of variance. A p≦0.05level of significance will be required for either test to reject theassumptions. If both assumptions are fulfilled, a single-factor ANOVAwill be applied, with animal grouping as the factor, utilizing a p≦0.05level of significance. If the parametric ANOVA is significant at p≦0.05,Dunnett's test will be used to identify statistically significantdifferences between the control group and each test article-treatedgroup at the 0.05 level of significance. If either of the parametricassumptions is not satisfied, then the Kruskal-Wallis nonparametricANOVA procedure will be used to evaluate intergroup differences(p≦0.05). The Dunn's multiple comparison test will be applied if thisANOVA is significant, again utilizing a significance level of p≦0.05.

Example 12 In Vivo Implantation of a Neo-Bladder Augmentation Constructafter Trigone-Sparing Cystectomy

Following implantation of neo-bladder augmentation constructs intocanine subjects as described in the Example above, the implantedconstructs were examined by fluoroscopic imaging, as well as forcapacity and compliance. FIG. 75 shows cystograms for the implantedconstructs at 4 months. A corresponds to a construct seeded withbladder-derived SMCs. B corresponds to a construct seeded withblood-derived SMCs; C corresponds to a construct seeded with adiposetissue-derived SMCs. D corresponds to native bladder baseline. FIG. 76shows the (A) capacity and (B) compliance of the implanted neo-bladderconstructs. All hematology and serum chemistries were found to be withinnormal limits for all bladder groups.

At 5 months, the animals appeared to be doing well clinically, gainingweight as expected, and all hematology and serum chemistries were foundto be within normal limits for all bladder groups. In addition, nohydroureter/hydronephrosis was observed. The adipose-derived SMC groupappeared to (i) have a higher mean bladder capacity versus theblood-derived SMC group, and (ii) be similar to the bladder-derived SMCgroup (10/10, 0.01/20 and 0/20) at 5 months. The compliance was similarbetween all groups at 5 months. FIG. 77 shows the average body weight ofthe animals. FIG. 78 shows the average serum creatinine of the animals.FIG. 79 shows the average BUN for the animals. FIG. 80 shows the averagealkaline phosphatase (ALP) for the animals. FIG. 81 shows the totalprotein average for the animals. FIG. 82 shows the white blood cellcount (WBC) average for the animals. FIG. 83 (blood) and FIG. 84(adipose) shows cystograms for the implanted constructs. Table 12.3shows the capacity of a native bladder versus a neo-bladder scaffoldconstruct.

TABLE 12.3 Native Scaffold Native Scaffold Native Scaffold Month5 >0.50 >0.65 >0.85 Bladder 10, 5 5 4 5 4 4 10 (N = 6) Bladder 0.1, 2 32 3 2 1 20 (N = 4) Bladder 0.01, 6 5 5 4 5 4 20 (N = 6) Bladder 0, 20 43 4 3 3 3 (N = 5) Blood 0, 20 5 5 4 3 4 2 (N = 6) Adipose 0, 20 5 4 4 33 2 (N = 6)

Example 13 Characteristics of a Total Regenerated Urinary Bladder

Introduction and Objectives: Neo-bladder durability and functionalityand the effect of total number smooth muscle cells (SMC) seeded onto aPLGA-based biodegradable polymer scaffold were evaluated in caninesfollowing radical cystectomy and implant of Autologous Neo-BladderReplacement constructs (NBR).

Methods: NBR were seeded with SMC at 3 densities: 25, 12, and 4×10⁶cells/Construct (n=8/grp). A group (n=8) where radically cystectomizedbladders were immediately reimplanted (R) served as a control. In-lifeassessments (radiographic, urinalysis, and urodyanamics) were performedfor the 9 mo study duration. Ex-vivo pharmacological and histologicalstudies were conducted on neo-bladder tissues at study termination.

Results: Animals were clinically healthy, continent, and able to urinateby 3 weeks post-implantation. At 9 mo post-implantation, all groups (NBRand R) had functional bladders with urodynamic compliance values andneo-bladder tissue histology (including mucosal and serosal linings,detrusor muscle, vasculature, and nerve components) consistent withnative bladder. Contractile responses to various concentrations ofcarbachol (Car) and phenylephrine (PE) were similar among all groups.However, contractile responses to α-β-methylene-ATP (AA) were evidentonly with R and NBR implants seeded with 25×10⁶ SMC. Logistic analysisof bladder tissue strips subjected to electrical field stimulation (EFS)revealed similar EC₅₀ and slope factor values for all groups.

Conclusions: An Autologous Neo-bladder Replacement Construct is capableof regenerating urinary bladder as a total organ that has structural,urodynamic, and pharmacological features similar to native bladder andis durable up to 9 mo after surgical implantation. There was no evidenceof abnormal tissue development, immune response, or evidence of systemicresponse to the neo-bladder regeneration. Pharmacological and urodynamicresponses suggest a positive correlation between number of autologousSMCs seeded per Neo-Bladder Replacement Construct and final regenerativeoutcome with 12×10⁶ SMCs achieving tissue regeneration and urodynamicoutcomes and 25×10⁶ SMCs achieving tissue regeneration, urodynamic, andpharmacologic outcomes similar to R.

Example 14 Adaptive Regulation of Regenerated Bladder Size

Homeostatic regulation that maintains organ size and structure is acomplex relationship between the specific organ, tissues, and bodyweight or size. Regulative development, or restoration of organ size andstructure after cell or tissue loss, can be observed during tissueregeneration or organogenesis. Some goals for regenerative therapiesinclude both restoration of structure and function and establishment ofadaptive regulation specific for the recipient. Adaptive regulation incystectomized animals implanted with cell-seeded PLGA-based scaffoldswas compared with adaptive regulation based on early results from aPhase II clinical trial of the Tengion Autologous NEO-BLADDER AUGMENT™(NBA) in children with neurogenic bladder due to spina bifida.

Neo-bladder capacity and body weight were measured in cystectomizedanimals implanted with cell-seeded PLGA-based scaffolds for 6 monthspost-implantation (p.i.). Cystometric capacity and voiding intervals(VI) were measured and formula predicted bladder capacity (FPBC) wascalculated at baseline and 12 months after NBA implant in two age- andweight-matched Phase II NBA clinical trial subjects (PT1 and PT2).

Implanted animals remained healthy and continent for study duration andachieved neo-bladder capacities consistent with body weight as early as6 months p.i. Histology and immunohistochemistry of neo-bladder tissuerevealed a native bladder-like structure and function, indicative ofbladder regeneration. PT1's baseline capacity was 33% of FPBC. At 12months p.i., PT1's capacity had increased 84% from baseline and achieved60% of FPBC. PT2 had capacities of 100% of FPBC at baseline and 12months p.i. VI increased for both PT1 and PT2.

These results demonstrate that autologous neo-bladders regenerated andgrew appropriately to the recipient's body size in animals and humans.These data support the conclusion that neo-bladders elicited by TengionAutologous NEO-BLADDER AUGMENT™ implantation are bioresponsive to theneeds of the recipient.

Example 15 Role of Biomechanical Stimulation (Cycling) in Neo-BladderRegeneration

Biomechanical stimulation is a process known to promote tissueregeneration and optimal healing. Bladder regeneration isbiomechanically stimulated by cycling (filling, storage and evacuation),a process that begins in utero and contributes to the development of afunctional bladder in humans by early childhood. Interruption of cyclingin patients with neurogenic bladder from either congenital (i.e., spinabifida) or acquired (i.e. spinal cord injury) impairment leads tosignificant functional and structural alternations.

Cycling impacts on bladder tissue regeneration in cystectomized animalsimplanted with cell-seeded PLGA-based scaffolds were evaluated andlearnings applied to outcomes of a Phase II clinical trial of theTengion Autologous NEO-BLADDER AUGMENT™ (NBA) in patients withneurogenic bladder due to spina bifida.

Post-implantation (p.i.) neo-bladder cycling was initiated in animals at2 weeks p.i. for 3 days/week. Three cycling parameters were collected:total weeks, hr/day, and total hrs. Urodynamic assessments from threecycling cohorts based on mean parameters were evaluated: HIGH (10weeks, >3.75 hr/day, >60 hrs), LOW (10 weeks, <2.25 hr/day, <25 hrs),and NO cycling. The HIGH cohort developed neo-bladders with improvedcompliance and capacities that were on average 3-fold higher than theLOW cohort (p<0.0001). The HIGH cohort achieved 90% of native baselinecapacity by 6 mo p.i., while the LOW cohort regained only 40%. Animalsnot cycled (incontinent) developed tubularized urinary tissuediversions. Histology of neo-bladder wall revealed more native-liketissue structure and extracellular matrix composition (e.g., elastin) incycled bladders. Early Phase II data studying the NBA suggest thatpatients with challenges in postoperative cycling (e.g. open bladdernecks, low pressure high grade reflux) had inferior clinical andurodynamic outcomes to patients without those challenges.

Early post-implantation cycling is essential for promoting regenerativehealing following implantation of autologous cell-seeded PLGA-basedscaffolds in animals and humans. Insights from Preclinical studies areconsistent with early insights from Phase II NBA trial and confirm theimportance of cycling in bladder regeneration.

FIG. 85 demonstrates the role of cycling in human urinary bladderdevelopment. Muscle and elastic fibers were found to increase whilecollagen was reduced. Sphincter tone develops near term to facilitatecycling dynamics (Wahl et al. BJU Int., 2003. 91:255). FIG. 86demonstrates that bladder capacity increases with age and urine output.An increase in urine production drives increased bladder capacity (Kimet al. J. Urol., 1991; 146:524).

FIG. 87A-C demonstrates that cycling influences regenerative outcome.Two canine groups with similar sized neo-bladder scaffolds were examinedand it was found that increased cycling resulted in higher capacitybladders. FIG. 87A depicts a histological comparison between animplanted neo-bladder that had been cycled versus an implantedneo-bladder that had not been cycled. Elastin fibers were observable inthe cycled bladder. FIG. 87B depicts the difference in capacity betweentwo different bladders based on the amount of time cycled. FIG. 87Cdepicts bladder capacity of a cycled bladder versus a non-cycledbladder.

FIG. 88 depicts the translation of regeneration-enhancing effects ofcycling to clinical outcomes. Human patients that had received aneo-bladder implant and were able to undergo cycling were observed tohave improved bladder capacity as compared to patients who were unableto undergo cycling. This suggests that cycling or biomechanicalstimulation promotes regeneration and is important for improvingclinical outcomes.

Example 16 Muscle Equivalent Constructs

Three-dimensional (3-D) constructs were fabricated from PGA/PLGA feltmaterial. Specifically, a porous degradable scaffold, constructed of PGAfelt coated with PLGA, was formed into a 3-D bladder-like shape, andseeded with cells. FIGS. 9A-E depict additional constructs.

In order to determine optimal scaffold formations, seven differentscaffold constructs were pre-formed and surgically tested forpreference. Qualitative scores were assigned to each construct using ascale of 1 to 10 (1=least liked; 10=most liked and preferred). Thescaffold constructs consisted of the seven scaffold constructs outlinedin Table 16.1 below.

TABLE 16.1 Scaffold constructs. Scaffold No. Scaffold Name Dimensions &Characteristics* 1 Pre-rolled, regular-sized, flat scaffold Ellipsoid 10cm long × 3.7 cm wide (see top conformation of FIG. 5a); 2D surface areaof 29.1 cm² Pre-rolled conformation is depicted in FIGS. 6c and 6d 2 2of 2 pre-rolled scaffold Two scaffolds, same length as scaffold No. 1,each of half width of scaffold No. 1, totaling the same surface area(see FIG. 8d) Pre-rolled conformation 3 Regular-sized scaffold Ellipsoid10 cm long × 3.7 cm wide (see top conformation of FIG. 5a); 2D surfacearea of 29.1 cm² 4 Piece of scaffold sterilized at 50° C. Sterilized ata higher temperature, which may result in more degradation and morepliability. 5 2 of 2 regular-sized scaffold Two scaffolds, same lengthas scaffold No. 1, each of half width of scaffold No. 1, totaling thesame surface area (see FIG. 8d) 6 Regular-sized scaffold cut in halfSame shape as scaffold No. 1, cut down center and sutured, in a mannersimilar to FIG. 8 7 Rectangular sheet from which to cut Rectangle forsurgeon to cut down as surgeon desired shape saw fit. *Except whereindicated otherwise, all scaffolds were sterilized at 30° C.

The results of the surgical preference testing are provided below inTable 16.2.

TABLE 16.2 Surgical Preference Test Results Scaffold Number Rating(1-10) Comments 1 6 Pre rolled ok will be difficult to lay flat 2 2 Toostiff will be difficult to manipulate 3 8 Good stiffness, easy tomanipulate  4* 8 Good stiffness, easy to manipulate 5 10 Easy to handle,good memory 6 9 Easy to handle, good memory, feels thin 7 1 Too stiffdifficult to handle, creases easy *Sterilized at a higher temperaturethan the other scaffolds.

Example 17 In Vivo

Study Schedule—in vivo: Pre-Study Procedures (Biopsy): Day (−20)-(−30);Study Start (Day 0): Approximately 20-30 Days Post Pre-Study Biopsy;Necropsy Start: Day 30±3 d and Day 84±3 d; Preliminary Report: 2-3 weeksafter receipt of pathology report; Final Report Issued: 2 weeks afterapproval of preliminary report; In-Vivo Study Completion: Day 84±3 d

Study Animal: Common Name: Yorkshire Pig; Breed/Class: Sus Scrofa;Number of Animals (by gender): 12 Female minimum; 2 Male minimum; AgeRange: On File; Weight Range: >45 kg

Study Design: The study design is shown below in Table 17.1 ((a): TissueDonor and Practice Animal (b): Scaffold mesh of poly(lactic-co-glycolicacid)+Autologous Smooth Muscle Cells (c) Urine Only (d) Pre-necropsysample for 30d animals (e) Smooth Muscle Cells=SMC).

TABLE 17.1 Study Design Number Blood & of Test SMC Vascular Urine Group# Animals Device Source Source Collection Cystogram Biopsy ImplantNecropsy 1 2 male Source^(a) NA NA NA NA ~20 NA NA days pre- implant 2 2female Construct^(b) Bladder Omentum Baseline; Baseline, NA Day 0 1F DayPeriodically Post-Sx, 30 +/− 3 for first 48 Wks 4^(d) & 1F Day hours^(c)8, Pre- 84 +/− 3 Wks 1, 2, 3, Necropsy 4^(d) & 8; Pre-Necropsy 3 2female Construct^(b) Bladder Peritoneum Baseline; Baseline, NA Day 0 1FDay Periodically Post-Sx, 30 +/− 3 for first 48 Wks 4^(d) & 1F Dayhours^(c) 8, Pre- 84 +/− 3 Wks 1, 2, 3, Necropsy 4^(d) & 8; Pre-Necropsy

Test Device: Neo-bladder Enlargement Construct w/ Bladder Smooth MuscleCells (SMC); Description: Scaffold No. 1, as described above in theExample above (Ellipsoid 10 cm long×3.7 cm wide (see top conformation ofFIG. 5A); 2D surface area of 29.1 cm²), comprising of syntheticlactide/coglycolide acid polymers plus autologous bladder smooth musclecells. Label Concentration: SMC number seeded on construct were providedon certificate of analysis. Storage Temperature: 22° C.±5.

Technical and Analytical Procedures

Pre-Surgery Fasting:

Procedure Description: All animals were fasted at least 8 hours prior tosurgery. Food was withheld the day prior to each surgery.

Duration/Frequency of the Procedure: Single event the night prior toeach surgery with duration of approximately 8 hours.

Body Weight:

Procedure Description: The weight of the animal was performed on acalibrated balance.

Duration/Frequency of the Procedure: Baseline, prior to implant surgery,weekly for the first month, then monthly thereafter (±2 days) andpre-necropsy.

Sedation/Anesthesia:

Procedure Description: Each animal was sedated and then anesthetizedprior to surgery preparation. Each animal was sedated with Ketamine 20mg/kg IM, and Xylazine 2 mg/kg (IM). Each animal was intubated andreceived inhalant isoflurane at 2.5%-4% for induction and 0.5-2.5% formaintenance of anesthesia, delivered through either a volume-regulatedrespirator or rebreathing apparatus. An intravenous sheath was placed ina peripheral vessel or right internal jugular vein. Lactated Ringer'ssolution was administered at 10 ml/kg/hr for the duration of thesurgical procedures (augmentation animals only).

Duration/Frequency of the Procedure: Throughout each day of in-lifeprocedures or surgery.

Surgical Preparation:

Procedure Description: For all animals (biopsy and augmentation), thehair over the entire abdominal region was clipped. The animal waspositioned in dorsal recumbency. A sterile urinary catheter was gentlyinserted into the bladder and the bladder emptied of urine prior to thestart of procedure.

Care was taken to ensure the bladder was not traumatized during theevacuation procedure. The operative area was then cleaned with threealternating scrubs of povidone-iodine solution and 70% alcohol; once thealternating scrubs were complete, a final application of povidone-iodinesolution was applied and allowed to dry. The area(s) was draped foraseptic surgery.

Duration/Frequency of the Procedure: Single Event Prior to Surgery

Biopsy (source male porcine only): Procedure Description: A midlineincision was made in the abdomen to allow access to the bladder. Priorto collecting the bladder tissue, the bladder was emptied of urine andone 2 cm×2 cm (approximate) piece of urinary bladder tissue was excised.

The biopsy tissues were immediately preserved aseptically in jarscontaining tissue culture media and then packaged. Additionally, >35 mLof venous or arterial blood was collected aseptically into an EtOsterilized plastic jar fortified with 0.05% heparin of total volume ofblood collected.

Duration/Frequency of the Procedure: Once per source animal lastingapproximately 1 hour per animal.

Catheter Implantation (Female Porcine Only):

Procedure Description: An indwelling catheter was placed within thejugular vein and within the bladder to facilitate blood and urinecollection in each animal.

Urinary Bladder Catheterization: A midline incision was made in theabdomen to allow access to the bladder. The bladder was emptied of urineand one 8-9.5 Fr open lumen catheter was inserted and sutured into thebladder to prevent movement. The insertion point was on the dorsal sideof the bladder away from the expected ventral enlargement site. Oncesecured to the bladder, the catheter was tunneled to the flank of theanimal where the port was attached and implanted in a subcutaneouspocket.

Jugular Vein Catheterization: The area surrounding the right or leftjugular vein was shaved and aseptically prepared. A 9.5 Fr siliconcatheter was inserted into the vein and secured by suture to preventmovement. Once secured, an extra large DaVINCI port was attached andimplanted in a subcutaneous pocket.

Duration/Frequency of the Procedure: Performed a minimum of 10 daysprior to enlargement surgery. The procedure will be performed once peranimal, approximately 1 hour per animal.

Urine Sample Collection and Analysis (Female Porcine Only):

Procedure Description: Two urine samples were collected bycatheterization or a pan caught method. Approximately 1.0 mL and 3.0 mLsamples were collected in sterile containers for qualitative andquantitative analysis, respectively. Smaller amounts were sufficient foranalysis; however, if urine collection was difficult, the quantitativesample took precedent over the qualitative sample. The collectedquantitative urine was decanted into 5 mL sterile tubes, refrigeratedand shipped within 24 hrs of collection. The qualitative measurement wastaken at time of collection using Multistix® 10 SG Test Strips.Parameters of interest for both the qualitative and quantitativemeasurements include: Glucose, Bilirubin, Blood, pH, Protein, Ketones,Urobilinogen, Specific Gravity, Nitrites, Bacteria 1 (quantitativeonly), and Leukocytes.

Duration/Frequency of the Procedure: Baseline, periodically for first 48hours, Weeks 1, 2, 3, 42 & 8 following implantation and prior tonecropsy. Procedure lasted approximately 15 minutes per animal.

Cystogram:

Procedure Description: The bladder of each animal was prepared forfluoroscopic imaging by: Placing a sterilized open lumen Foley catheterinto the bladder access site; • Attaching a syringe and drawing outurine to empty the bladder; Injecting 3:1 saline: contrast medium viathe open lumen catheter and filling the bladder; Performing fluoroscopicimaging.

Duration/Frequency of the Procedure: Baseline, post-surgery, Week 42, 8and pre necropsy. Procedure lasted approximately 10-30 minutes peranimal.

Blood Collection (Female Porcine Only):

Hematology, Coagulation, Serum Chemistry and Blood Gases Hematology:Hematology samples were collected in 2.0 ml EDTA tubes, and stored onwet ice or refrigerated (2-8° C.). Samples were labeled and packaged onice. Analysis was performed within 24 hrs of collection. Blood sampleswere evaluated for the parameters specified below: Total leukocyte count(WBC); Erythrocyte count (RBC); Hemoglobin concentration (HGB);Hematocrit value (HCT)a; Mean corpuscular volume (MCV); Mean corpuscularhemoglobin (MCH)a; Mean corpuscular hemoglobin concentration (MCHC)a;Platelet count (PLT); Reticulocyte count (RTC); White blood celldifferential; a=Calculated values.

Coagulation: A total of 1.8 mL blood was collected into 1.8 mL sodiumcitrate tubes (0.2 mL of 3.8% sodium citrate). Citrated blood sampleswere kept on ice until ready to be centrifuged at 1,700×g for 15minutes. Before freezing, plasma was divided in half and freezon at −70°C. One vial was sent to designated laboratory and the other was kept asa back-up until end of study. Citrated plasma samples were stored at−70° C.

Coagulation parameters measured include: Prothrombin time (PT);Activated partial thromboplastin time (APTT); • Fibrinogen (FIB).

Serum Chemistry: Samples for Serum Chemistry were collected inapproximately 4.0 ml serum separation tubes. The blood samples werecentrifuged (10,000 RPM for 10 minutes) and the serum extracted usingsterile technique. Serum was evenly split among two separate labeledvials. Serum was frozen at −70° C. Serum samples were evaluated for thefollowing parameters: Glucose (GLU); Urea nitrogen (BUN); Creatinine(CRE); Total protein (TPR); Albumin (ALB); Globulin (GLOB) 1;Albumin/Globulin ratio (A/G) 1; Calcium (CAL); Phosphorus (PHOS);Electrolytes: Sodium (NA), Potassium (K), and Chloride (CL); Totalcholesterol (CHOL); Total bilirubin (TBIL); Triglycerides (TRG); Alanineaminotransferase (ALT); Aspartate aminotransferase (AST); Alkalinephosphatase (ALK); Gamma glutamyltransferase (GGT); 1=Calculated values

Blood Gases: Blood Gas/Spun Hematocrit/Total Protein Monitoring:Arterial blood samples (˜1.0 mL) were collected and analyzed using acalibrated i-STAT analyzer and the appropriate cartridge. Samples wereevaluated for the following blood gas parameters: Sodium (Na) (mmol/L);Potassium (K) (mmol/L); Ionized Calcium (iCa) (mmol/L); Glucose (Glu)(mg/dL); Hematocrit (Hct) (%); pH, PCO2 (mm Hg); PO2 (mm Hg); TCO2(mmol/L); HCO3 (mmol/L); BEecf (mmol/L); pH; and SO2(%).

Duration/Frequency of the Procedure Collection of blood samples wereconducted as follows: Baseline, Weeks 1, 2, 3, 4 (pre-necropsy samplefor 30 day animals) & 8 following implantation and prior to necropsy.Procedure lasted approximately 15 minutes per animal.

Vital Signs Monitoring Procedure: Procedure Description: The followingvitals were monitored during implant surgery:

Body Temperature: Body temperature was monitored through an esophagealprobe.

Direct Blood Pressure and Heart Rate: A blood pressure line wasconnected to monitor systolic arterial pressure (SAP), diastolicarterial pressure (DAP), and mean arterial pressure (MAP). Anintroducing sheath was placed in the carotid artery for blood pressuremonitoring.

Expired gases: CO2 and SaO2

ECG: Lead II ECG was monitored

Duration/Frequency of the Procedure: At approximately 20 minuteintervals throughout the surgical procedure.

Laparoscopic Bladder Enlargement:

Procedure Description:

Positioning and Port Placement: A four port transperitoneal techniquewas employed to gain laparoscopic access to the bladder andintraperitonal space. Four punctures were made: a 12-mm primary port wasinserted ˜1 cm above the umbilicus; two 12-mm secondary ports wereinserted ˜7-10 cm lateral to and ˜3-4 cm below the umbilicus; a 5-mmsuprapubic port was inserted to establish pneumoperitoneum using CO²and/or facilitate laparoscopic handling of the omentum, peritoneum andmanipulation of neo-bladder. Alternatively, a fifth abdominal puncturebelow the umbilicus was performed using a Veress needle to establishpneumoperitoneum using CO2 until the pressure reaches approximately 15mm Hg.

Bladder Enlargement: Three bladder enlargement techniques were performeddepending on the vascular source being utilized (i.e., omentum orperitoneum).

Bladder Enlargement using Peritoneum: A suitable segment of peritoneumable to exceed the distance to the primary 12-mm port without tensionwas carefully isolated from the abdominal wall intracorporeally. Asegment of peritoneum larger than the size of the construct was thenexternalized through one of the 12-mm ports and the peritoneum carefullyspread out to accommodate the construct. The seeded construct was thenremoved from the media and the construct number matched to the animaldocumentation for verification. The construct was then secured to theperitoneum via surgical adhesive or suture allowing peritoneum tooverlap the entire construct. The construct was maintained moist duringthe procedure using a sterile syringe and gently infusing sterilephysiological pH saline. Once the construct was secured, theperitoneum/construct unit was carefully internalized through a 12-mmport into the intraperitonal space and positioned longitudinally on thebladder dome to just above the urethra on the ventral side of thebladder (opposite to the ureter orifices). One side of the construct wastacked to the bladder using appropriately sized staples, e.g., 0.45 cmhorizontal dimension×0.47 cm vertical dimension. Once securely attachedto the bladder, a longitudinal incision was made into the bladdermimicking the position of the construct and the incised bladder tissuestapled to the non secured side of the construct. Preventive measureswere taken to limit the amount of residual urine within the incisedbladder from reaching the abdominal cavity. Any peritoneum overlappingthe construct was then secured to the bladder using surgical adhesive.

Bladder Enlargement using Omentum: A suitable segment of terminalomentum able to reach the entire length of the bladder without tensionwas gently grasped using endoscopic clamps and carefully isolated fromthe abdominal cavity intracorporeally. The omental segment waspositioned longitudinally on the bladder dome to just above the urethraon the ventral side of the bladder (opposite to the ureter orifices).One side of the omentum vascular was tacked to the bladder surface usingsurgical adhesive. Once the omentum segment was securely attached to thebladder, the seeded construct was then removed from the media and theconstruct number matched to the animal documentation for verification.The construct was carefully internalized through a 12-mm port into theintraperitonal space and positioned longitudinally on the bladdermimicking the secured omentum line. The construct side nearer thesecured omentum was tacked longitudinally onto the bladder usingappropriately sized staples. A longitudinal incision was made into thebladder along the same line as the secured construct. The incisedbladder tissue was stapled to the non secured side of the construct.Preventive measures were taken to limit the amount of residual urinewithin the incised bladder from reaching the abdominal cavity. Anyomentum overlapping the construct was then secured to the bladder usingsurgical adhesive. Using a lumen catheter, the augmented bladder waschecked for leaks to assure adequate closure and water-tightness. The 4laparoscopic ports were removed and the abdominal punctures were thenclosed with absorbable suture material of an appropriate size. The skinwas closed in a subcuticular fashion, using an appropriate size ofabsorbable suture material.

Duration/Frequency of the Procedure: Once per animal lastingapproximately 5 hours per animal on Day 0.

Bladder Enlargement using Omentum via Laparotomy: The omentum bladderenlargement procedure outlined above was altered from laparoscopy tolaparotomy. For comparison purposes, two peritoneum bladder enlargementprocedures were performed as a laparotomy, as shown below in Table 17.2.

TABLE 17.2 Laparotomy Study Design Necropsy Post- DB-Pig Scaffold CellOrigin & # Tissue Wrap Implant Time 1 Scaffold Pig Bladder- Omentum 30 ±2 days No. 1* 2 derived SMCs Omentum 84 ± 5 days 3 (10 × 10⁶) Peritoneum30 ± 2 days 4 Peritoneum 84 ± 5 days *as described above

Briefly, a midline incision was made in the abdomen beginningimmediately caudal to the umbilicus. The omentum and peritoneum wereidentified, carefully separated from the abdominal space until thetissue is long enough to cover the enlarged bladder section. Care wastaken to ensure the tissue remained vascularized. The urinary bladderwas then exposed and the bladder carefully emptied of urine taking careto avoid urine from entering into the abdominal cavity. The augmentationof the construct to the bladder followed the same procedure outlinedabove. Once secured, the abdominal incision was closed in layers withabsorbable suture material of an appropriate size. The skin was closedin a subcuticular fashion, using an appropriate size of absorbablesuture material.

Duration of the Procedure: Once per animal lasting ˜5 hours per animalon Day 0.

Post-Surgery and Recovery Analgesia:

Procedure Description: Survival animals receive the following:Antibiotic Therapy—Approximately 2 mg/kg Naxcel (ceftiofur) orequivalent antibiotic was provided intramuscularly to each animal priorto and following enlargement surgery beginning on Day 0. Treatmentcontinued once daily until Day 3 or until the facility veterinariandeemed appropriate.

Postoperative Analgesia—Approximately 0.1 mg/kg, IM given 8-12 hoursapart, of Buprenorphine was administered following surgery (enlargementprocedure [Day 0]) for a total of two injections. Analgesic therapycontinued twice a day for up to 3 days following the definitive surgery(Days 1-3) for a total of six injections.

Concurrent Therapy—As prescribed by the Facility Veterinarian and StudyDirector, animals were provided concurrent therapy (e.g., antibiotics orfluid therapy) in order to maintain general good health.Duration/Frequency of the Procedure: Immediately following surgicalprocedures and daily thereafter at the discretion of the FacilityVeterinarian.

Animal Sacrifice and Necropsy

Procedure Description:

Unscheduled and Scheduled Euthanasia—All animals were injected withsodium pentobarbital (150 mg/kg, IV) to cause euthanasia. Scheduledeuthanasia was day of biopsy (male porcine only), Day 30 and Day 84post-enlargement procedure.

Necropsy—All female animals will be subjected to necropsy. There was aspecific focus on the urinary bladder. The complete bladder (trigone,anastomotic site, and neo-bladder) was visualized and photographed insitu and then excised en bloc and fixed in 10% NBF.

Duration/Frequency of the Procedure: Single event duration ofapproximately ½ hr per animal on Day 30 & 84 (±3 days).

Histology/Pathology Laboratory

Procedure Description: The fixed urinary bladder (i.e., enlargedneobladder) was trimmed to include separate sections across theinterface between the normal bladder and the construct. Tissue sampleswere trimmed, embedded in paraffin, and sectioned. Slides will bestained with hematoxylin and eosin (H & E) and Masson's Trichrome(elastin).

Duration/Frequency of the Procedure: Histology and Pathology wereconducted within 3 months of receipt of samples.

Results: As shown in Tables 17.3 and 17.4 below, the implantation didnot affect the animals' capacity for growth as measured by body weightand the associated increase in the bladder's volumetric capacity. FIG. 8f shows a cystogram of the implanted patch of the instant invention at 4weeks.

TABLE 17.3 Body Weight. Body Weight (Kg) Animal Number Implant 28 Days55 Days 1 60.0 71.7 — 2 55.9 67.8 81.6 3 63.1 71.2 89.7 4 56.8 68.0 —

TABLE 17.4 Bladder Capacity. Bladder Capacity (mL) Animal Number Implant28 Days 55 Days 1 700 1200 — 2 750 1100 2350 3 775 1250 2650 4 1100 1400—

FIG. 89 shows the implantation of a scaffold.

FIG. 90 shows a cystogram of an implanted patch scaffold at 4 weeks.

Example 18 Adipose-Derived Smooth Muscle Cells Versus Mesenchymal StemCells (MSCs)

Adipose tissue represents a heterogenous cell population composed ofendothelial cells, adipocytes, smooth muscle cells and progenitor cellswith limited mesenchymal differentiation potential. We have usedquantitative RT-PCR, antigen expression, protein fingerprinting, growthkinetics and functional analysis to evaluate the cellular composition ofthe adherent, stromal vascular fraction (SVF) derived from humanadipose. We show that enrichment for the smooth muscle cell compartmentof adipose SVF is directly dependant on media formulation. These humanadipose-derived smooth muscle cells (Ad-SMC) are functionallyindistinguishable from human bladder-derived smooth muscle cells andphenotypically and functionally distinct from mesenchymal stem cells(MSC) or other adipose-derived progenitor populations.

We have investigated the cellular composition of the initial “passagezero” adherent human SVF-derived cell population using quantitativereal-time PCR methods (TaqMan). We show that though this startingadherent SVF-derived cell population is composed of cells expressingendothelial, smooth muscle and adipocyte-associated markers, we havebeen able to identify and culture a cell population with markedlydistinctive biological properties through the expansion of SVF-derivedcells under defined media conditions that select against the growth ofMSC (Gong et al. Tissue Eng Part A 2008; 15:319-330; Lund et al.Cytotherapy 2009; 11:189-197). Despite partial overlap indifferentiation potential and expression of markers historicallyassociated with MSC, this cell population clearly has a more pronouncedsmooth muscle cell phenotype relative to MSC based on FACS and RT-PCR(reverse transcription PCR) analysis of the expression of key nuclearand cell surface markers. This population also expresses noticeablyfewer endothelial-specific genes when compared to MSC. Manifestation ofa smooth muscle cell phenotype is independent of passage number, adiposedonor source or the requirement for directed differentiation withrecombinant cytokines and growth factors. Additionally, this smoothmuscle cell enriched population has a distinctive proteomic signaturewhich unambiguously discriminates it from MSC. Finally, we haveleveraged the diametrically opposing responses of this smooth musclecell like population and MSC towards the thromboxane A2 mimetic U46619to document a clear functional dichotomy between the two cell types.Taken together, these data support the conclusion that this populationis more accurately described as adipose-derived smooth muscle cells(Ad-SMC), and represents a separate and distinctive cellular speciescompared to other classes of adipose-derived cells including adipocytes,endothelial cells and MSC.

Methods and Materials.

Preparation of Adipose Tissue. Human adipose samples were obtainedeither subcutaneously or through lipoaspiration (Zen-Bio, ResearchTriangle Park, N.C.), and washed 3-5 times with an equal volume ofPBS/gentamycin (Gibco) (5 μg/ml). Adipose was digested withfilter-sterilized collagenase I (Worthington) (0.1%, 1% BSA in DMEM-HG(Gibco)) at 37° C. for 1 hour, then centrifuged for 5 minutes at 300 gin 50 ml conical tubes. The stromal vascular fraction was resuspended inPBS/1% BSA and filtered through a 100 μm Steriflip vacuum filter. Thecell population was pelleted again at 300 g for 5 minutes andresuspended in DMEM-HG+10% FBS+gentamycin 5 μg/ml. Bone marrow derivedMSC at the end of passage two was obtained from a commercial supplier(Lonza). For studies on the effect of media type on expression of smoothmuscle cell markers, the SVF-derived cells were alternativelyresuspended in α-MEM (Gibco)+10% FBS, SMCM (ScienCell) or L15 (Sigma).

Taq-Man qRT-PCR. RNA was purified from MSC or Ad-SMC using the RNeasyPlus Mini Kit (Qiagen) according to the manufacturer's instructions.cDNA was generated from 2 μg of RNA using the SuperScript VILO cDNASynthesis Kit (Invitrogen) according to the manufacturer's instructions.Following cDNA synthesis, each sample was diluted 1:10. qRT-PCR wassetup as follows using the TaqMan primers and probes listed below: 10 μlmaster mix (2×), 1 μl primer/probe, 9 μl cDNA (diluted 1:10).

The following TaqMan primers were used for evaluation of smooth muscle,endothelial and adipogenic gene expression: SmαA (smooth muscle alphaactin): Hs00909449_m1, SM22: Hs00162558_m1, myocardin: Hs00538076_m1,SMMHC (smooth muscle myosin heavy chain): Hs00224610_m1, calponin:Hs00154543_m1, adiponectin: Hs00605917_m1, FABP-4 (fatty acid bindingprotein #4): Hs1086177_m1, CDH5/VECAD (vascular endothelial cadherin):Hs00174344_m1, vWF (von Willebrand factor): Hs00169795_m1, PECAM1(platelet endothelial cell adhesion molecule #1): Hs00169777_m1,FLT1/VEGFR (VEGF receptor): Hs01052936_m1, KDR/FLK1 (fetal liver kinase#1): Hs00176676_m1, TEK (tyrosine kinase, endothelial): Hs00945155_m1.18 s rRNA was used as endogenous control and all samples were calibratedagainst bladder smooth muscle cell cDNA. All primer/probes were securedfrom Applied Biosystems. All reactions were carried out in an ABI 7300real time thermal cycler using default cycling parameters. Analysis ofPCR data was performed using the method of Relative Quantitation (RQ) byComparative Ct.

Array-RT-PCR. Real time array-based qRT-PCR analysis was performed for35 cycles using the SABiosciences MSC (PAHS-082A) and Cell SurfaceMarker PCR array platform (PAHS-055A) according to the manufacturer'sinstructions.

FACs analysis. 0.5×10⁶-1×10⁶ cells per data point were fixed in 2%paraformaldehyde and F_(c) receptors blocked to prevent non-specificbinding. Cells were then incubated with a directly conjugated antibodyfor the cell surface markers CD31, CD45, CD54, CD56, CD73, CD90, CD105,CD117 or CD133 (BD Biosciences) as recommended by the manufacturer.Subsequent to final washing (PBS, 0.1% Triton X-100), antigen detectionwas performed utilizing the BD FACS Aria 1 or Guava EasyCyte MiniExpress Assay system using the appropriate fluorescent channel. Aminimum of 5000-10,000 events were acquired from each sample.

2D Proteomic analysis. Passage controlled (end of P2) bone marrowderived MSC and Ad-SMC were lysed in Lysis Buffer (50 mM Tris pH 8; 150mM NaCl; 0.5% NP40 and protease inhibitors, Roche) and 40 μg of proteinlysate from each cell type was run out on a pH 4.0-7.0 Zoom IEF strip(Invitrogen) according to the manufacturer's instructions. Each stripwas then loaded onto a 4-12% Bis/Tris acrylamide gel and run out on the2^(nd) dimension. The gels were stained with SYPRO Ruby stain(Invitrogen) according to the manufacturer's instructions.

Passage controlled (end of P2) bone marrow derived MSC and Ad-SMC werelysed in Lysis Buffer (50 mM Tris pH 8; 150 mM NaCl; 0.5% NP40 andprotease inhibitors, Roche) and 40 μg of protein lysate from each celltype was run out on a pH 4.0-7.0 Zoom IEF strip (Invitrogen) accordingto the manufacturer's instructions. Each strip was then loaded onto a4-12% Bis/Tris acrylamide gel and run out on the 2^(nd) dimension. Thegels were stained with SYPRO Ruby stain (Invitrogen) according to themanufacturer's instructions.

Results

Expression markers in Ad-SVF. We performed a quantitative TaqMan RT-PCRanalysis of the cell population derived from the stromal-vascularfraction of adipose tissue adherent on the tissue culture flask withinthe initial 24-48 hours subsequent to plating, using a panel of definedendothelial, adipocytic and smooth muscle cell specific TaqMan primers.This served to analyze expression markers in the initial adherent cellpopulation as well as establishing a baseline for subsequent analysis ofthe effects of passage, time and media formulation upon expression ofsmooth muscle cell specific genes. As shown in FIG. 91A, low butdetectable levels of FABP-4 and adiponectin were observed in theadherent cell population within the first 24 hours, consistent with thepresence of residual adipocytes. Similarly, an endothelial populationdefined by expression of VECAD, vWF, PECAM, FLT1, FLK and TEK waspresent at this time point (FIG. 91D-E). A smooth muscle cell populationdefined by expression of SMαA, SM22, myocardin, SMMHC and calponin wasalso observed within the earliest adherent cell population (FIG. 91B-C).We were able to detect all three cell populations at comparable levelswithin 24-48 hrs of plating. As discussed below, smooth muscle cellswere isolated from this mixture of cell populations.

Expression of smooth muscle markers is dependent on media type. Asadipose is a heterogenous tissue composed of multiple cell types, it isreasonable to expect that enrichment for smooth muscle cells overendothelial cells or MSCs may be affected by media formulation.Isolation of undifferentiated MSCs from bone-marrow and adipose isclosely dependant on media composition (Gong et al. 2009 supra). Inparticular, the presence of elevated levels of glucose in the media orgrowth at high density appears to select against the expansion of MSC(Lund et al. 2009 supra; Stolzing et al. Rejuv Res 2006; 9:31-35). Wereasoned that modulation of media formulation may be useful inenrichment for smooth muscle cells at the expense of MSC and other cellpopulations. As shown in FIG. 92A-B (Taqman analysis of SMC markerexpression by media type), the expansion of a smooth muscle cellenriched population from adipose-SVF is tightly dependent upon growth inDMEM-HG media. Expansion in α-MEM, SMCM or L15 is associated with amarkedly reduced smooth muscle cell phenotype as shown by decreasedexpression of SMαA, SM22, myocardin, SMMHC and calponin.

Ad-SMC more closely resemble smooth muscle cells than MSC. We usedsemi-quantitative RT-PCR to assess the smooth muscle cell associatedgene expression signatures of Ad-SMC and MSC. As shown in FIG. 93, theexpression of the key smooth muscle markers calponin, myocardin andSMMHC is noticeably more pronounced in Ad-SMC when compared to MSC,supporting our hypothesis that this cell population is more similar tosmooth muscle cells than to MSC. We then evaluated the stability ofexpression of SMC specific markers across multiple independent adiposepreparations (n=4) and over 5 passages in culture. As shown in FIG. 94(RT-PCR of Ads across passage), the expression of SMαA, SM22, SMMHC,myocardin and calponin is remarkably constant across passage and isindependent of donor, demonstrating that expression of a smooth musclecell phenotype is stable over time. These observations are consistentwith Ad-SMC being a more fully differentiated, phenotypically stablecell population.

Array-based RT-PCR analysis demonstrates significant differences in geneexpression of key markers between Ad-SMC and MSC. We have used theSABiosciences MSC Marker Array panel to systematically identifydifferences in gene expression between passage controlled (P2) Ad-SMCand MSC. This panel profiles the expression status of 84 genes involvedin MSC pluripotency and self-renewal. A summary of the key markersidentified as distinct between Ad-SMC and MSC is shown in Table 18.1.

TABLE 18.1 Fold Regulation Hu Ad-SMC vs Symbol Description MSC Ct Ad-SMCP4 Ct MSC BMP6 Bone morphogenetic protein 6 34.09 28.57 35.5555 CD44CD44 molecule (Indian blood group) 31.87 23.06 347.7725 IL1B Interleukin1, beta 35 29.62 32.2673 GDF5 Growth differentiation factor 5 25.3431.89 −120.9276 HGF Hepatocyte growth factor (hepapoietin A; scatter27.95 32.77 −36.4539 factor) LIF Leukemia inhibitory factor (cholinergicdifferentiation 29.37 35 −63.9113 factor) MCAM Melanoma cell adhesionmolecule 27.99 33.67 −66.1652 RUNX2 Runt related transcription factor 227.69 30.58 −12.1426 VCAM1 Vascular cell adhesion molecule 1 24.36 34.1−1103.5987 Historically Defined Cell Surface Markers Fold RegulationSymbol Description MSC Ct Hu Ads P4 Ct MSC vs. Ads ALCAM Activatedleukocyte cell adhesion molecule (CD166) 24.88 24.44 1.0512 ENG Endoglin(CD105) 23.06 22.57 1.0882 NT5E 5′-nucleotidase, ecto (CD73) 25.2 24.381.3679 THY1 Thy-1 cell surface antigen (CD90) 29.54 29.68 −1.4221

Significant (at least ten fold) down-regulation in Ad-SMC relative toMSC was observed for GDF5, HGF, LIF, MCAM, RUNX2 and VCAM1. Significant(at least ten fold) up-regulation in Ad-SMC compared to MSC was observedfor BMP6, CD44, and IL1β. These key differences in gene expression wereobserved to remain consistent independent of passage or cell sample(n=6, data not shown).

Gene expression analysis was continued using the SABiosciences SurfaceMarker Array. A summary of the key results is presented in Table 18.2,where we have examined Ad-SMC at P0 and P4.

TABLE 18.2 PCR Array Catalog # PAHS-055A Human Cell Surface Markers onHuman Adipose Zhu05 SQ RT 300xg DMEM Cell Type Symbol Description Ct P0Ct P4 Fold Regulation P0 to P4 SMC MYH9 Myosin, heavy chain 9,non-muscle 23.59 24.2 −2.5245 MYH10 Myosin, heavy chain 10, non-muscle25.94 26.05 −1.7851 MYOCD Myocardin 35 33.09 2.2721 Endothelial ENGEndoglin (Osler-Rendu-Weber syndrome 1) 23.84 22.73 1.305 ICAM2Intercellular adhesion molecule 2 27.35 29.02 −5.2634 NOS3 Nitric oxidesynthase 3 (endothelial cell) 30.01 31.66 −5.191 PECAM1*Platelet/endothelial cell adhesion molecule (CD31 antigen) 29.07 35−100.8453 SELP Selectin P (granule membrane protein 140 kDa, antigenCD62) 35 35 N/A TEK* TEK tyrosine kinase, endothelial (venousmalformations, 31.27 25.68 29.1212 multiple cutaneous and mucosal)VCAM1* Vascular cell adhesion molecule 1 25.88 34.16 −514.1338 VWF* VonWillebrand factor 27.2 31.49 −32.3569 Adipocyte RETN Resistin 35 35 N/AFibroblast ALCAM Activated leukocyte cell adhesion molecule 28.6 24.987.4333 COL1A1 Collagen, type I, alpha 1 20.26 20.65 −2.1675 COL1A2Collagen, type I, alpha 2 19.36 18.2 1.351 HLA HLA-A Majorhistocompatibility complex, class I, A 24.53 24.7 −1.8609 HLA-DRA* Majorhistocompatibility complex, class II, DR alpha 26.57 34.12 −309.9733CD74* CD74 molecule, major histocompatibility complex, 28.13 35−193.4746 class II invariant chain Other NT5E 5′-nucleotidase, ecto(CD73) 27.43 24.24 5.5174 NCAM1 Neural cell adhesion molecule 1 35 35N/A *= Change in Fold Regulation > |10.0|

Expression of the fibroblastic/stromal markers ALCAM, COL1A1 and COL1A2is maintained across passage, as are the smooth muscle cell specificmarkers MYH10, MYH9 and MYOCD. The population is negative for theadipocyte marker RETN, indicating that there is minimal contaminationwith adherent adipocytes. Importantly, although Ad-SMC acquire an HLAMHC II negative status within 4 passages they are initially HLA MHC IIpositive, a key distinction with MSC which are MHC II negative. Anotherinteresting observation is that Ad-SMC becomes progressively lessendothelial with passage, as judged by the general trend indown-regulation of the endothelial markers ENG, ICAM2, NOS3, PECAM1,SELP, TEK, VECAM and VWF. This data is independently confirmed by theRT-PCR analysis in FIG. 95.

To further compare the gene expression profiles between theadipose-derived smooth muscle cells (Ad-SMC) and mesenchymal stem cells(MSC), a PCR-based gene array analysis was performed for humanmesenchymal stem cell markers (SABiosciences; PCR Array Catalog#PAHS-082A) (data not shown). The results illustrated the extent ofhomologous gene expression among Ad-SMC, MSC, and a well characterizednon-MSC cell type, human aortic endothelial cells (HuAEC). Of the 84human MSC genes analyzed, human Ad-SMC share only 27% homology (23 of 84genes) with human MSC at initial isolation (data not shown). Incontrast, the well characterized, non-MSC, HuAEC share 49% homology (41of 84 genes) with MSC (data not shown). This supports the conclusionthat the Ad-SMC share significantly less homology with MSC than HuAEC,which is a well-known non-MSC cell type. Thus, the Ad-SMC are even lesslike MSCs than HuAEC are, further supporting the conclusion that theAd-SMC cells isolated from adipose tissue are Ad-SMC and not MSC.

The cell surface profile of Ad-SMC is significantly different from thatdefined for MSC. We observed that both MSC and Ad-SMC share expressionof the surface markers CD73, CD90, CD105 and CD166 which aretraditionally associated with MSC (Table 18.1). However, as discussedbelow, these markers have no intrinsic biological significance beyondtheir historical association with MSC. The gene expression results fromthe cell surface marker RT-PCR analysis were generally reflected in thecomparative FACs analysis presented in FIG. 96A-C (Ad-SMCs) and FIG.97A-B (MSCs), which shows that Ad-SMC are CD31+, CD45+, CD54+, CD56+,CD90+, CD105+. Importantly, Ad-SMC was CD45+ and CD117+, a cleardistinction from MSC, which are CD45− CD117−. Expression of CD73 isconsistent with that previously reported for adipose stromal vascularfraction (da Silva Meirelles et al. J Cell Sci., 119:2204 (2006)), butdiffers from that reported for bone-marrow derived MSC. We were alsoable to observe a small but distinct population of CD133+ cells,possibly reflecting a small, pluripotent cell sub-population.

Passage controlled MSC and Ad-SMC have unique proteomic signatures. FIG.98 shows a comparative analysis of the whole proteomic signatures ofMSC, bladder-derived SMC, Ad-SMC, and human aortic smooth muscle cells.The top two panels demonstrate that Ad-SMCs are distinct from MSC andare also clearly different from MSC isolated from adipose tissue as wellas other classes of stem and progenitor cells (Roche et al; Proteomics2009; 9:223-232; Noel et al. Exp Cell Res 2008; 314:1575-1584). Thearrows on both gels highlight one difference between MSCs and AdSMCs;concentration of proteins at different and distinct locations within thepH gradient and molecular weight range. MSC have this proteinconcentration closer to a pH of 7.0, and greater than or equal to 60,000molecular weight. In contrast, AdSMC have this protein concentrationcloser to a pH of 4.0, and less than 60,000 in molecular weight. AdSMCalso had more protein present with pI above 7 than MSC, as indicated bythe smear along the right outside edge of the gel at pH 7.0. Bladdersmooth muscle cells were analyzed as a control. The boxes indicate areasof similarity among all samples. It is clear that the AdSMC proteinprofile is most like the profile for bladder-derived SMC (lower leftpanel), which is distinct from the pattern observed for MSC. Aorticsmooth muscle cells were also analyzed as an additional smooth musclecell control (lower right panel). The proteomic signature of the aorticsmooth muscle cells and bladder smooth muscle cells are almostidentical. Taken together, the high degree of similarity among theprofiles for AdSMC, bladder and aortic smooth muscle cells, which aredistinctly different from the profile of the MSC, supports theconclusion that SMCs, not MSCs, are being isolated from adipose tissue.All gels were stained with SPRYO Ruby stain to visualize the proteinpattern.

Growth kinetics of Ad-SMC differ markedly from MSC. The proliferativepotential of Ad-SMC differs markedly from MSC which have beensuccessfully expanded to up to 40 passages (Bruder et al., J CellBiochem., 64:278-294 (1997)). As shown in FIG. 99, Ad-SMC show a markeddecline in proliferative capacity after the 4th-5th day in culture. Wehave also observed that unlike MSC, Ad-SMC exhibit contact dependantinhibition of proliferation. These observations demonstrate that Ad-SMChave no capacity for self-renewal and therefore by definition are notstem or progenitor cells. MSCs do not exhibit contact inhibition ofproliferation and they can be observed piling on top of each other,similar to foci formation in transformed cell cultures. This isconsistent with previous observations (Zhou et al. 2006 supra).

Ad-SMC and MSC have distinctly opposing responses to treatment withU46619. As part of our efforts to evaluate the effects of smallmolecules targeting signaling cascades involved in the activation ofsmooth muscle cell related developmental pathways, we have focused onU46619, a thromboxane A2 mimetic whose effects include increasingintracellular Ca²⁺ levels and activation of RhoA, CaM and MLC kinasesignaling cascades. As reported previously (Kim et al. 2009, Stem Cells.27(1):191-199), we have confirmed that treatment with U46619 (1 μM) ledto up-regulation of the key smooth muscle cell markers myocardin andSMMHC in MSC. However, Ad-SMC responded to the same treatment byunambiguous downregulation of myocardin and SMMHC expression as shown inFIG. 100. These results provide clear evidence for a functionaldichotomy between Ad-SMC and MSC.

Expression of functional markers. FIG. 101 provides results of RT-PCRanalysis of mesodermal differentiation markers. Lane contents: 1: MSCcontrol; 2: MSC experimental; 3: AdSMC control; 4: AdMSC experimental;5: Peripheral blood control; 6: Peripheral blood experimental; and 7:H₂O. The expression of markers of mesodermal differentiation in MSC andAdSMCs undergoing adipogenic differentiation. AdSMC shows significantlygreater expression of ostepontin relative to MSC during growth understandard conditions (n=1). Expression of Oct4B, a splice-variant ofOct4A, an established marker for pluripotentiality (Kotoula et al.,2008, Stem Cells 26(1): 290-1), is significantly upregulated inadipose-derived cells relative to MSC. Neither MSC nor adipose-derivedcells show expression of Oct4A.

FIG. 102 shows the results of RT-PCR analysis of Oct4A/Oct4B expressionin MSC/AdSMC. Lane contents: 1: Bladder SMC; 2: HFF-1 (humanfibroblast); 3: MSC; 4: AdSMC; 5: peripheral blood; 6: H₂O. Expressionof the closely related transcriptional isoforms Oct4A and Oct4B wasevaluated in MSC, AdSMC, fibroblast and SMC lines. No expression of thepluripotency marker Oct4A (Gong et al. 2009 supra) was observed, thoughall cell lines evaluated expressed Oct4B (n=1).

Discussion. In this report, we have evaluated the marker expression ofadipose-derived cell populations. Adipose tissue represents aheterogenous mix of cell types, including endothelial cells, pericytes,smooth muscle cells, adipocytes and MSC (Lin et al., Stem Cells Dev2008; 17:1053-1063). Adherent cells from the stromal vascular fractionof adipose isolated under different conditions and expanded at differentdensities and media formulations are often grouped together as MSC, withno systematic approach towards establishing cellular composition acrossdisparate conditions (Rebelatto et al., 2008 supra; Liu et al. 2007supra; Jack et al. Biomaterials 2009; 30:3259-3270). Similarly, adherentcells from the mononuclear fraction of bone marrow are typicallyreferred to collectively as MSC. However, multiple laboratories claim tohave isolated distinct bone-marrow derived stem cell or progenitorpopulations with disparate but overlapping phenotypic and functionalproperties, though it remains to be determined whether or not theserepresent unique cell types in vivo (Ulloa-Montoya et al., J BiosciBioeng 2005; 100:12-27; Ratajczak et al. Folia Histochem et Cyto 2004;42:139-146; Lodie et al. Tissue Eng 2002; 8:739-751). Likewise, a numberof studies have reported conflicting conclusions regarding the degree offunctional and phenotypic similarity between adipose and bone-marrowderived MSC (Roche et al. Proteomics 2009; 9:223-232; Noel et al. ExpCell Res 2008; 314:1575-1584). The identification of any non-bone marrowderived stromal cell population as MSC has been questioned through theapplication of in vivo heterotopic ossicle formation assays, whichindicate that only bone marrow derived stromal cells may be labeled asMSC (Kalz et al. Stem Cells 2008; 26:2419-24). These datanotwithstanding, it may be reasonably concluded from evaluation of thepublished literature that adipose-derived MSC and bone-marrow derivedMSC share overlapping but distinctive differentiation potentials, asevaluated by quantitative PCR-based lineage analysis (Roche et al. 2009supra; Noel et al. 2008 supra; Rebelatto et al. 2008 supra; Liu et al.2007 supra).

We began our analysis of the initial, adherent adipose stromal vascularfraction-derived cells using TaqMan Q-RTPCR. As shown in FIGS. 91 & 92,we were able to isolate a cell population that consistently displays asmooth muscle cell phenotype as shown by expression of SMαA, SM22,SMMHC, calponin and myocardin. Although endothelial and adipocyticmarkers are also detectable within the initial 24-48 hour windowsubsequent to plating, the population retains smooth musclecharacteristics as the incidence of other cell characteristics decreasewith passage (FIGS. 91, 92, Table 7). Expression of adipogenic markerswas observed to rapidly decline (Table 7). Unlike other reportsdescribing the isolation of MSCs from adipose and their subsequentapplications in tissue engineering, no inductive cytokines oradditional, exogenous growth factors are required to directdifferentiation of a smooth muscle associated gene expression signature(Jack et al. 2009 supra).

Ad-SMCs are directly comparable to bladder-derived smooth muscle cellsas defined by gene and protein expression of smooth muscle cellassociated markers and Ca²⁺ dependant contractility (Basu et al., 2009in preparation; Basu et al. International Society for Stem CellResearch, 7^(th) Annual Meeting, Jul. 8-11, 2009). Isolation of Ad-SMCsis directly dependant upon isolation in a particular mediaformulation—as shown in FIG. 91B, expression of smooth muscle cellmarkers is contingent upon expansion in DMEM-HG media. Growth in othermedia types leads to loss of smooth muscle cell associatedcharacteristics and possibly leads to enrichment for more mesenchymalprogenitor populations (Gong et al. 2009 supra). To this end, thepresence of high glucose levels in media or expansion at high densityhas been shown to be detrimental to the manifestation of a robust MSCdifferentiation potential (Lund et al. 2009 supra). Additional studieshave shown increased osteogenic differentiation potential of MSC in lowglucose media compared to high glucose media (Jager et al. Biomed Tech(Berl) 2003; 48:241-244). It has been suggested that the presence ofadvanced glycation end products related to glucose and other sugars maylead to loss of differentiation potential in MSC (Kume et al. J BoneMiner Res 2005; 20:1647-1658). Taken together, these observationsdemonstrate that expansion of adipose-SVF derived cells under conditionsof high density and high glucose leads to selection for a smooth musclecell phenotype and against acquisition of MSC characteristics.

Continuing with the gene expression approach, we have used the Array PCRdata panel in Tables 18.1 and 18.2 to identify a core group of markersthat consistently and unambiguously discriminates Ad-SMC from MSC. BMP6,CD44 and IL-1β show at least 30 fold greater expression in Ad-SMCcompared to MSC, whereas GDF5, HGF, LIF, MCAM, RUNX2 and VCAM1demonstrate at least 30 fold greater expression in MSC relative toAd-SMC. These results are consistent across multiple donor samples(n=3), suggesting that our observations are not a consequence of donorvariability or random fluctuations in gene expression levels. BMP6 is amember of the TGF-β superfamily which has been implicated in theregulation of chrondrogenesis and osteogenesis during MSCdifferentiation (Henning et al. J Cell Physiol 2007; 211:682-291;Friedman et al. J Cell Biochem 2006; 98:538-554). Our data showingup-regulation of BMP6 expression clearly discriminates Ad-SMC fromadipose-derived MSC where down-regulation of BMP6 expression wasobserved relative to bone marrow derived MSC (Henning et al. 2007supra). Interestingly, induction of adipose-derived MSC with exogenousBMP6 led to an up-regulation in expression of the TGF-β1 receptor(Henning et al. 2007 supra). The TGF-β signaling pathway is wellestablished to have a critical role in activation of smooth muscle cellspecific developmental pathways Owens et al. Acta Physiol Scand 1998;164:623-635. CD44 is a well known marker of MSC-like cells and has beenwidely implicated in cell growth, migration and homing (Khaldoyanidi S.Cell Stem Cell 2008; 2:198-200). Expression of CD44 between adipose andbone-marrow derived MSCs has been shown to be similar in terms ofoverall expression, regulation of transcriptional splice variants andthe overall stability of gene expression (Peroni et al. Exp Cell Res2008; 314:603-615).

Genes found to be up-regulated in MSC over Ad-SMC include GDF5, whichhas been shown to be important in the regulation of chondrogenic andosteogenic differentiation in MSC, and HGF. HGF and its cognate receptorc-met have been identified with the regulation of motility andproliferation in bone-marrow derived MSC (Neuss et al. Stem Cells 2004;22:405-414). Consistent with earlier reports, we have observedexpression of the pro-inflammatory cytokine LIF to be a keydistinguishing feature separating MSCs and Ad-SMCs (Majumdar et al. JHematother Stem Cell Res 2000; 9:841-8). Importantly, LIF acts as a keymarker of progenitor status in MSC, serving as a proxy for maximumdifferentiation potential (Whitney et al. Tissue Eng Part A 2009; 15:1).This observation is in agreement with our interpretation that Ad-SMCrepresents a smooth muscle cell population. MCAM (CD146) is a cellsurface marker closely associated with MSC derived from the perivascularniche of adipose (Zannettino et al. J Cell Physiol 2008; 214:413-421)and bone-marrow (Baksh et al. Stem Cells 2007; 25:1384-92). Theexpression of CD146 appears to be closely correlated with the stem cellpotential of MSC-like cell populations from adipose or bone-marrow(Zannettino et al. 2008 supra; Baksh et al. 2007 supra; Gronthos et al.J Cell Physiol 2001; 189:54-63). RUNX2 is a transcription factorinvolved in regulation of osteogenesis during differentiation of MSC(Isenmann et al., Stem Cells 2009). Expression of the cell-adhesionmarker VCAM1 (CD106) is also characteristic of MSC isolated from adipose(Zannettino et al. 2008 supra) or bone-marrow (Brooke et al. Stem CellsDev 2008; 17:929-40). Finally, Ad-SMC show strong expression of MHCClass II, unlike MSC isolated from either adipose or bone marrow(Niemeyer et al. Tissue Eng 2007; 13:111-121).

Taken together, the gene expression data suggest that Ad-SMC represent amore fully differentiated SMC population, rather than an MSC-like cellpopulation. This interpretation is corroborated by the 2D whole proteomecomparison of Ad-SMC with MSC shown in FIG. 39, which demonstrates thatAd-SMC and MSC have distinctive and unique proteomic signatures.Additional comparison of the Ad-MSC proteomic profile with that reportedfor adipose-derived MSC and other classes of stem or progenitor cellshows little, if any, significant overlap (Noel et al. 2008 supra;Rebelatto et al. 2008 supra). Given the associated expression ofmultiple mature smooth muscle cell markers and loss of endothelialmarkers as well as functional contractility comparable tobladder-derived smooth muscle cells (Basu et al., 2009; in preparation;Basu et al. International Society for Stem Cell Research, 7^(th) AnnualMeeting, Jul. 8-11, 2009), these data strongly suggest that Ad-SMC arein fact smooth muscle cells rather than MSCs.

In parallel with the gene expression studies discussed above, we haveexamined the expression of key MSC-associated cell surface markers onboth bone-marrow derived MSC and Ad-SMC by FACS. Both cell types wereconsistently positive for CD90+ and CD105+ but were negative for CD73, awell established marker for MSC, suggesting the potential forconsiderable heterogeneity in the expression of standard MSC markers(Chamberlain et al. Stem Cells 2007; 25:2739-49). Furthermore, Ad-SMCwere observed to be CD45+ CD117+, (FIG. 96) which unambiguouslydiscriminates them from MSC derived from either adipose or bone marrowsources (Lee et al. Cell Physiol Biochem 2004; 14:311-324).Identification of a CD45+ compartment suggests the existence of asub-population of hematopoeitic origin, unlike MSC. These observationsnotwithstanding, we believe that the identification of cell surfacemarkers such as CD73, CD90 and CD105 with MSC has no intrinsicbiological significance, and may be viewed as an artifact created duringthe historical progression of the field (Dominici et al. 2006 supra).Although AD-SMC may share some of these archetypal cell surface markerswith MSCs (for example, CD90 and CD105), they are clearly distinct fromMSC in the expression of other established markers (such as CD34, CD45and CD117). It is therefore challenging to believe that these markershave discriminatory value, as we and others have observed multiple,fully differentiated cell types to robustly express many of the samemarkers commonly associated with MSC (Jones et al. Rheumatology 2008;47:126-131). The combined transcriptomic, proteomic and functionalanalysis of MSC and Ad-SMC presented in the current report will likelybe more useful in evaluating whether or not MSC and Ad-SMC representbiologically distinct cell populations (Lodie et al. 2002 supra; Gong etal. 2009 supra).

Our functional comparison of Ad-SMC and MSC focused on the analysis ofgrowth kinetics, smooth muscle phenotype and response to small moleculedrugs targeting smooth muscle cell specific signaling pathways. A keyfeature of stem cells is the capacity for self-renewal. MSC have thecapacity for self-renewal as demonstrated by the ability to expand forat least 25-40 passages while retaining the potential for multi-lineagedifferentiation (Tintut et al. Circulation 2003; 108:2505-2510; Reyes etal. Blood 2001; 98:2615-2625; Bruder et al. J Cell Biochem 1997;64:278-294). In contrast, as shown in FIG. 99, Ad-SMC demonstrate asharp drop in growth potential within 4-5 days initial plating,consistent with their identification as a terminally differentiatedsmooth muscle cell type. There is no indication of any capacity forself-renewal.

Another characteristic of stem and progenitor cell populations is therequirement for directed differentiation along defined developmentallineages using a combination of exogenous growth factors, ECM and othercontrollable components of the extracellular milieu. A number of reportshave focused on the regulated differentiation of MSC from adipose orbone marrow for applications in tissue engineering and regenerativemedicine. For example, adipose-derived MSC were differentiated intosmooth muscle like cells using inductive media containing 100 U/mlheparin for up to 6 weeks prior to seeding polymeric bladder dome-likescaffold structures that demonstrated evidence of functionality in a ratcystectomy model (Jack et al. 2009 supra). In addition, TGF-β or smallmolecule agonists targeting the TGF-β signaling pathway includingsphingosylphosphorylcholine, bradykinin and angiotensin II have alsobeen used for induction of a smooth muscle like phenotype from adiposeor bone marrow derived MSC (Gong et al. 2009 supra; Kim et al. CellSignal 2008; 20:1882-1889; Jeon et al. 2006 supra; Kim et al. Int JBiochem Cell Biol 40; 2482-2491). A less targeted approach, epigenomicreprogramming with the DNA demethylating agent 5-azaC, has been used todirect bone marrow-derived MSC towards a cardiomyocyte-like phenotype(Xu et al. Exp Biol Med 2004; 229:623-631). Dedifferentiated adipocytesmay also be driven along a smooth muscle lineage using TGF-β and havebeen reported to contribute towards bladder tissue regeneration in amouse bladder injury model (Sakuma et al. J Urol 2009 July;182(1):355-65. Epub 2009 May 20). Finally, methods for TGF-β induceddifferentiation of smooth muscle cells from bone-marrow derived cellshave been described (Kanematsu et al. Am J Pathol 2005; 166:565-573;Becker et al. Int J Artif Organs 2008; 31:951-9). Taken together, thesereports typically present a MSC-like population with little or noexpression of any smooth muscle cell associated markers prior totreatment with an inductive cytokine or small molecule agonist.

In marked contrast, we have been able to directly isolate and expand asmooth muscle cell population from adipose expressing all key smoothmuscle associated markers including those typically associated withmature smooth muscle cells (SmαA, SM22, SMMHC, calponin and myocardin)without the requirement for directed differentiation by any exogenousagent (Owens et al. Physiol Rev 2004; 84:767-801). This observationstrongly suggests that Ad-SMC represent a cell population that isalready more fully differentiated from initial isolation andfundamentally distinct from MSC. The acquisition of smooth muscle likefeatures by porcine bone-marrow derived MSC after multiple passaging athigh density without the addition of exogenous growth factors has beenrecently presented (Shukla et al. World J Urol 2008; 26:341-349).However, in contrast to this report, we are able to isolate Ad-SMC witha clear smooth muscle cell phenotype from the earliest passage acrossmultiple, independent preparations, (n=174 as demonstrated in FIG. 94).There is no requirement for “differentiation” through prolonged growthat confluence.

Co-ordinate regulation of multiple smooth muscle cell specific geneexpression pathways with signaling cascades regulating contractilitythrough disparate independent kinases, so-called“excitation-transcription” coupling (Wamhoff et al. Circ Res 2006;98:868-878) is modulated by U46619, a stable analog of thromboxane A2(TxA2). Treatment with U46619 has been shown to lead to increasedexpression of SRF and myocardin in adipose-derived MSC and associatedup-regulation of the smooth muscle cell specific markers SMαA, calponin,smoothelin and SMMHC (Kim et al. 2009 supra). The effect of TxA2 onexcitation-transcription coupling appears to serve as a functionalfingerprint for MSC, regardless of the tissue of origin. We haveobserved that bone-marrow derived MSC recapitulates the up-regulation ofsmooth muscle markers upon treatment with U46619 as observed foradipose-derived MSC (Kim et al. 2009 supra). However, as shown in FIG.100, Ad-SMC respond to U46619 in a diametrically opposing manner,showing unambiguous down-regulation of the key functional smooth musclemarkers myocardin and SMMHC. Clearly, the organization and regulation ofsignaling cascades involved in excitation-transcriptional coupling asobserved in MSC is fundamentally different in Ad-SMC. This observationprovides definitive evidence that Ad-SMC is functionally distinct fromadipose or bone-marrow derived MSC and in fact represents a biologicallyunique cell population.

From where do Ad-SMCs originate and what is their relationship to MSC?Adipose is a heavily vascularized tissue and a number of studies haveimplicated the perivascular niche as a potential source of both MSC aswell as smooth muscle and endothelial cells (Caplan J Pathol 2009;217:318-324). Pericytes with MSC differentiation potential have beenisolated directly from blood vessels as well as from multiple organsystems throughout the body (da Silva Meirelles et al. 2006 supra; daSilva Meirelles et al. Tissue Eng Part A 2009 February; 15(2):221-9;Tintut et al. 2003 supra). However, although SMαA+ cells have beenlocalized to all capillaries, arterioles and venules of theadipose-derived vascular bed, expression of STRO-1, a key MSC-specificmarker, is tightly associated with endothelium and additionally foundonly within a subset of blood vessels (Lin et al. Stem Cells Dev 2008;17:1053-1063). Furthermore, expression of the stem cell-specific markersOct4 and telomerase was observed only rarely, suggesting that trulypluripotent progenitors are uncommon within adipose (Lin et al. 2008supra). In their entirety, these observations point to MSC, endotheliumand smooth muscle occupying distinct spaces within the broaderperivascular niche. Nevertheless, there remains the potential forconsiderable ebb-and-flow across developmental lineages. For example,endothelial cells appear capable of lineage switching towards a smoothmuscle cell like phenotype in response to TGF-β or the depletion ofpro-angiogenic factors and loss of endothelial cell-cell contact(Krenning et al. Trends Cardiovasc Med 2008; 18:312-323; Krenning et al.Biomaterials 2008; 29:3703-3711).

Finally, adherent cell types with endothelial and smooth musclephenotypes as well as limited mesenchymal differentiation potential havebeen identified to circulate in adult peripheral blood (He et al. StemCells 2007; 25:69-77). Such circulating smooth muscle cells maycontribute to the population of adipose-derived smooth muscle cells,although we have been unable to purify them directly from human adultperipheral blood in meaningful numbers (our unpublished observations).Given that MSC in long term culture also follow a smooth muscle celllike differentiation pathway (Dennis et al. Stem Cells 2002;20:205-214), we believe that taken together, the published data as wellas our observations are consistent with the perivascular niche as asource for a broad continuum of smooth muscle, endothelial and MSC celltypes with variable degrees of proliferative and differentiationpotential. Although smooth muscle cells generated by the directeddifferentiation of MSC or isolated directly from adipose continue toshow evidence of mesenchymal differentiation plasticity (Kim et al. 2009supra; our unpublished observations), nonetheless, we believe a cleardistinction may be drawn between opposing ends of the spectrum, withadipose-derived smooth muscle cells having unambiguous functional andphenotypic differences relative to MSC of bone-marrow or adipose SVForigin.

In closing, we have demonstrated that the isolation of Ad-SMCs directlyfrom the P0 adherent stromal vascular fraction of adipose is tightlydependant on media formulation. Expression of smooth muscle cell markersis robust and consistent and is independent of donor source and acrosspassage. We have shown that Ad-SMCs are phenotypically distinct from MSCas demonstrated by gene expression, proteomic and surface markeranalysis, and are functionally distinct from MSC as evaluated by theirresponse to pharmacologic agents targeting smooth muscle cell associatedsignaling pathways. In contrast to other published reports, isolation ofthese smooth muscle cells does not require directed differentiation withTGF-β or related small molecules. Ad-SMC may be expanded to up to 10⁷cells within 4-5 passages, express the full range of smooth muscle cellassociated markers and are functionally comparable to bladder-derivedSMC both in vitro (Ca²⁺-dependant contractility) and in vivo(regeneration of neo-urinary conduit in swine cystectomy model) (Basu etal., 2009 in preparation; Basu et al. International Society for StemCell Research, 7^(th) Annual Meeting, Jul. 8-11, 2009). These datasupport the conclusion that this population is more accurately describedas adipose-derived smooth muscle cells (Ad-SMC), and represents aseparate and distinct population compared to other classes ofadipose-derived cells including endothelial cells and MSC.

1. A urinary diversion for a defective bladder in a subject comprisinga) a first implantable, biocompatible construct comprising a tubularscaffold having a first end configured to form a stoma and connect to anabdominal wall section, a second closed end, and at least a first sideopening configured to connect to a first ureter; and b) an autologouscell population that is not derived from the defective bladder,deposited on or in a surface of the scaffold, wherein the urinarydiversion is adapted to provide passage of urine out of the subject. 2.The urinary diversion of claim 1 wherein the scaffold further comprisesa second side opening configured to connect to a second ureter.
 3. Theurinary diversion of claim 1 wherein the first end is configured to bepositioned flush with the abdominal wall.
 4. The urinary diversion ofclaim 3 wherein the first end is configured to be sutured to the skin ofthe subject.
 5. The urinary diversion of claim 1 wherein the stomafurther comprises a stoma button.
 6. The urinary diversion of claim 1,wherein the biocompatible scaffold is biodegradable.
 7. The urinarydiversion of claim 1, wherein the scaffold comprises a material selectedfrom the group consisting of polyglycolic acid, polylactic acid, and acopolymer of polyglycolic acid and polylactic acid.
 8. The urinarydiversion of claim 1, wherein the cell population is a smooth musclecell population.
 9. The urinary diversion of claim 1, wherein thediversion is free of urothelial cells.
 10. The urinary diversion ofclaim 8, wherein the smooth muscle cell population is derived fromadipose.
 11. The urinary diversion of claim 8, wherein the smooth musclecell population is derived from peripheral blood.
 12. The urinarydiversion of any one of claims 8, 10, and 11, wherein the SMC populationis not derived from in vitro differentiation of mesenchymal stem cells(MSCs).
 13. The urinary diversion of claim 1, wherein the first end isan open end configured for anastomosis to an opening in the subject'sabdominal wall.
 14. The urinary diversion of claim 1, wherein the firstend is an open end configured to be anastomosed to the skin.
 15. Theurinary diversion of claim 1, wherein the tubular scaffold is coatedwith a biocompatible and biodegradable shape-setting material.
 16. Theurinary diversion of claim 15, wherein the shape-setting materialcomprises a poly-lactide-co-glycolide copolymer.